POTENTIAL APPLICATION OF VARIABLE SURFACE …
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POTENTIAL APPLICATION OF VARIABLE SURFACE GLYCOPROTEINS IN DIAGNOSIS OF TRYPANOSOMA BRUCEI RHODESIENSE SIMON NGAO MULE MASTER OF SCIENCE (Biotechnology) JOMO KENYATTA UNIVERSITY OF AGRICULTURE AND TECHNOLOGY 2016
Transcript of POTENTIAL APPLICATION OF VARIABLE SURFACE …
POTENTIAL APPLICATION OF VARIABLE SURFACE
GLYCOPROTEINS IN DIAGNOSIS OF TRYPANOSOMA
BRUCEI RHODESIENSE
SIMON NGAO MULE
MASTER OF SCIENCE
(Biotechnology)
JOMO KENYATTA UNIVERSITY OF
AGRICULTURE AND TECHNOLOGY
2016
Potential application of variable surface glycoproteins in diagnosis of
Trypanosoma brucei rhodesiense
Simon Ngao Mule
A thesis submitted in partial fulfilment for the degree of Master of
Science in Biotechnology in the Jomo Kenyatta University of
Agriculture and Technology
2016
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DECLARATION
This thesis is my original work and has not been presented for a degree in any other
university.
Signature..................................................Date……………………………
Simon Ngao Mule
This thesis has been submitted for examination with our approval as the university
supervisors
Signature…………………………………..Date………………………………
Dr. Juliette R. Ongus
JKUAT, Kenya
Signature…………………………………..Date………………………………
Dr. Vincent O. Adung’a
International Center of Insect Physiology and Ecology, Kenya
Signature…………………………………..Date……………………………….
Dr. Daniel K. Masiga
International Center of Insect Physiology and Ecology, Kenya
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DEDICATION
I dedicate this work to my parents, Mr. Joshua Mule and Mrs. Rosemary Mule for their
unwavering support, belief, guidance, devotion and prayers during my studies.
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ACKNOWLEDGEMENTS
I would like to acknowledge, foremost, the Almighty God for the strength, good health
and a sober mind during this study. I express my deepest gratitude to my supervisor, Dr.
Daniel Masiga, for the chance to undertake my masters project under his supervision and
his expertise which greatly assisted the research. I am greatly thankful to Dr. Vincent
Owino, who patiently corrected my manuscript, review and thesis drafts, and whose
comments greatly improved my writing skills. I thank Dr. Juliette Ongus for reviewing
my thesis and for the positive criticism.
This work was partly supported by the National Commission for Science, Technology and
Innovation (NACOSTI), the World Federation of Scientists (WFS) and icipe’s Dissertation
Research Internship Programme (DRIP). Without these organizations, this work would
not have been successfully accomplished.
I am grateful to my colleagues for the different input to this study.
Finally, I am greatly indebted to my parents, sisters Mukami and Mwende, my brother
Musyoka, my nieces Mutio and Mutheu, and my nephew Mumo for always supporting
me with prayers, advice, encouragement and best wishes.
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TABLE OF CONTENTS
DECLARATION .............................................................................................................. ii
DEDICATION ................................................................................................................. iii
ACKNOWLEDGEMENTS ............................................................................................ iv
LIST OF TABLES ............................................................................................................ ii
LIST OF FIGURES ........................................................................................................ iii
ABBREVIATIONS ......................................................................................................... iv
ABSTRACT ...................................................................................................................... v
CHAPTER ONE .............................................................................................................. 1
1.0 INTRODUCTION ...................................................................................................... 1
1.1 Background .................................................................................................................. 1
1.2 Statement of the problem ............................................................................................. 4
1.3 Justification .................................................................................................................. 5
1.4 Research Hypothesis .................................................................................................... 6
1.5 Objectives ..................................................................................................................... 6
1.5.1 General Objective...................................................................................................... 6
1.5.2 Specific Objectives.................................................................................................... 6
CHAPTER TWO ............................................................................................................. 7
2.0 LITERATURE REVIEW .......................................................................................... 7
2.1 African Trypanosomiasis .............................................................................................. 7
2.1.1 African Animal trypanosomiasis (AAT).................................................................... 7
2.1.2 Human African Trypanosomiasis (HAT) ................................................................ 11
2.2 Trypanosome life cycle .............................................................................................. 13
2.3 Control of African trypanosomiasis ........................................................................... 15
2.3.1 Chemotherapy ......................................................................................................... 16
2.3.2 Vector Control ......................................................................................................... 17
2.4. Diagnosis of Human African Trypanosomiasis......................................................... 20
2.4.1. Clinical Diagnosis .................................................................................................. 20
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2.4.2 Parasitological Diagnosis ........................................................................................ 21
2.4.3 Serological Diagnosis ............................................................................................. 23
2.4.4 Molecular Tools....................................................................................................... 30
2.5 Disease Stage Determination ..................................................................................... 33
2.6 Antigenic Variation in African Trypanosomes ........................................................... 36
2.7. The Variable Surface Glycoprotein (VSG) ............................................................... 38
2.8 Sequential Expression of VSG ................................................................................... 39
2.9. New serodiagnostic algorithms ................................................................................. 39
CHAPTER THREE ....................................................................................................... 41
3.0 MATERIALS AND METHODS ............................................................................. 41
3.1 Generation of Recombinant VATs in Bacterial and Insect Expression Systems ........ 41
3.1.1 Recovery of Cloned Variable Antigen Types (VATs) .............................................. 41
3.1.2 Propagation and Purification of Recombinant Plasmids. ........................................ 42
3.1.3 Cloning and Expression of VATs in Bacterial Cells ................................................ 42
3.1.4 Cloning and Expression of VATs in Insect Cells ..................................................... 49
3.2 Evaluation of the Diagnostic Potential of the Generated Recombinant VATs ........... 52
3.2.1 Infection of Monkey disease models and sera collection ....................................... 52
3.2.2 Serum and Immunodiffusion Gel Set-up ................................................................ 53
3.2.3 Evaluation of Diagnostic Potential via Ouchterlony Immuno-assays..................... 54
CHAPTER FOUR .......................................................................................................... 55
4.0 RESULTS .................................................................................................................. 55
4.1 Generation of Recombinant VATs in Bacterial and Insect Expression Systems ........ 55
4.1.1 Expression of Recombinant VATs in Bacterial Cells .............................................. 55
4.1.2 Expression of Recombinant VATs in Insect Cells ................................................... 57
4.2 Evaluation of the Diagnostic Potential of the Expressed Recombinant VATs ........... 61
4.2.1 Serodiagnostic Potential of Recombinant VAT3 and VAT4 .................................... 61
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4.2.2 Potential Application of VATs for Early Disease Detection .................................... 62
CHAPTER FIVE ............................................................................................................ 70
5.0 DISCUSSION ........................................................................................................... 70
5.1 Generation of Diagnostic Recombinant VATs in Bacterial and Insect Expression
systems…... ...................................................................................................................... 70
5.2 VATs Have Diagnostic Potential for Antibody Detection .......................................... 71
CHAPTER SIX .............................................................................................................. 74
6.0 CONCLUSIONS AND RECOMENDATIONS ..................................................... 74
6.1 CONCLUSIONS ........................................................................................................ 74
6.2 RECOMMENDATIONS ........................................................................................... 74
REFERENCES ............................................................................................................... 76
APPENDICES ................................................................................................................ 97
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LIST OF TABLES
Table 3-1: Serially diluted L-Arabinose for protein expression pilot studies .......... 46
Table 3.2: A table showing the different T. b. rhodesiense strains used for infecting
monkeys belonging to four experimental groups, and their years of isolation. ............... 53
Table 4-1: Diagnostic performance of recombinant VATs against monkey sera samples
infected with four different T. brucei rhodesiense KETRI strains collected at different days
post infection (dpi) ........................................................................................................... 64
Table A-1: Sequences of primers used to amplify VAT3 and VAT4 genes ............... 95
Table A-2: Quantification results of Purified recombinant VATs expressed in bacterial
and insect cells ................................................................................................................. 98
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LIST OF FIGURES
Figure 2-1: Tsetse fly and cattle distribution in Africa. ......................................... 8
Figure 2-2: Human African trypanosomiasis (HAT) distribution, incidence and risk for
travellers in sub Saharan Africa. ...................................................................................... 13
Figure 2-3: Life cycle of Trypanosoma brucei in mammalian host and the tsetse fly
vector……. ....................................................................................................................... 15
Figure 2-4: Lumber puncture to collect CSF for subsequent staging of sleeping
sickness….. ...................................................................................................................... 35
Figure 2-5: African trypanosome antigenic variation. .......................................... 37
Figure 4-1: Results of VAT3 and VAT4 amplification, cloning and bioinformatics
analysis…… ..................................................................................................................... 56
Figure 4-2: SDS-PAGE analysis of expression and purification of recombinant VAT3
and VAT4 in bacterial cells............................................................................................... 57
Figure 4-3: Agarose gel results of VAT3 and VAT4 gene amplification and
bioinformatics results of recombinant pIZ/V5-his constructs. ......................................... 58
Figure 4-4: Microscopy and SDS PAGE results of recombinant protein expression
in insect cells (Sf21 cell line). .......................................................................................... 60
Figure 4-5: Ouchterlony double diffusion results showing immune recognition of
recombinant VATs and sera collected from T. b. rhodesiense infected monkeys ............. 62
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ABBREVIATIONS
AAT African Animal Trypanosomiasis
CATT Card Agglutination Test for Trypanosomiasis
CIATT Card Indirect Agglutination Test
DNA Deoxyribonucleic Acid
ESAG Expression Site-Associated Gene
FAO Food and Administration Organization of the United Nations
FIND Foundation for Innovative New Diagnostics
GPI Glycosylphosphatidylinositol
HAT Human African Trypanosomiasis
IPTG Isopropyl-Beta-D-Thiogalactopyranoside
JKUAT Jomo Kenyatta University of Agriculture and Technology
LB Luria Bertani medium
PCR Polymerase Chain Reaction
SDS-PAGE Sodium Dodecyl Sulphate Polyacrylamide Gel Electrophoresis
TE Tris-EDTA buffer
TLF1 Trypanolytic Factor 1
TLF2 Trypanolytic Factor 2
TNM-FH Tricoplusia ni medium-formulation hink
VAT Variable Antigen Type
VSG Variable Surface Glycoprotein
WHO World Health Organization
X-gal 5-Bromo-4-Chloro-Indolyl-Β-D- Galactopyranoside
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ABSTRACT
Sleeping sickness, also known as human African trypanosomiasis (HAT), is a neglected
tropical disease caused by two subspecies of Trypanosoma brucei namely T. b. gambiense
and T. b. rhodesiense. Trypanosoma b. gambiense causes chronic form of HAT (Gambian
HAT) in West and Central Africa while T. b. rhodesiense is responsible for the acute form
(Rhodesian HAT) in East and Southern Africa. In both cases, the disease evolves through
clinically distinct stages namely early/stage 1 and late/stage 2. The early stage is
haemolymphatic, in which parasites develop in the blood, lymph and peripheral organs.
The parasites then spread to the central nervous system (CNS) (late/encephalitic stage)
where they cause serious neurological disorders, and results into death if untreated. This
necessitates timely diagnosis, stage determination and treatment, with drugs used
dependent on the stage of the disease. However, diagnostic tools applied are limiting,
suffering from low sensitivity and specificity, high costs and inapplicability in field
environments in rural areas where the disease is endemic. This calls for development of
improved and/or novel diagnostic tools, specifically for Rhodesian HAT, which kills in
weeks or months if not treated. One potential diagnostic approach is serodiagnosis using
trypanosome surface proteins, the variable surface glycoproteins (VSGs), which are
highly immunogenic and expressed in an ordered fashion early during infection. In this
study, two VSGs previously shown to be predominantly expressed early during T. b.
rhodesiense infection in monkeys were expressed in bacterial and insect expression
systems, and the purified recombinant diagnostic antigens used for immunological
detection by applying Ouchterlony double immunodiffusion test. The recombinant
proteins generated from both expression systems detected anti-VSG antibodies in a panel
of sera from infected monkeys, with a detection rate of 89%, 97% and 83% by VSG3
expressed in bacterial cells, VSG3 and VSG4 expressed in insect cells respectively. In
addition, sera recovered up to 15 days post infection, and 19 days when the parasite has
crossed into the CNS were seropositive, suggesting potential application of VSGs in
diagnosis of sleeping sickness. This is a proof of concept that VSG recombinant antigens
predominantly expressed during early stage of trypanosome infection have diagnostic
potential, and could form a basis for the development of a simple, cheap, robust and
sensitive screening tool applicable in the field setting for early disease detection.
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CHAPTER ONE
INTRODUCTION
1.1 Background
African trypanosomes (Genus: Trypanosoma) are single cell protozoan parasites that
cause sleeping sickness or human African trypanosomiasis (HAT) in man and Nagana or
animal African trypanosomiasis (AAT) in cattle; both diseases are generally called African
trypanosomiasis (AT). The parasite is transmitted by tsetse fly (Genus: Glossina), an insect
vector found in 37 countries in sub-Saharan Africa where the disease is endemic, with an
estimated 70 million people (WHO, 2012; Simarro et al., 2013) and 60 million cattle
(Cecchi et al., 2014) at different levels of infection risk. This region is over 10 million
square kilometres (Cecchi and Mattioli, 2009) and represents the most arable part of the
continent. HAT affects approximately 10, 000 people annually, with 3,796 new cases
reported by the World Health Organization (WHO) in 2014 (WHO, 2015), a new low in
75 years. This reported HAT incidence could however be an underestimate due to
inaccessibility of some of the most affected areas, poor or no health infrastructure, logistic
constrains and lack of accurate diagnostic tools (Cattand et al., 2001). AAT, on the other
hand, affects the health and productivity of livestock in approximately 348 distinct disease
loci (Cecchi et al., 2014), and results in 3 million cattle deaths annually across the tsetse
infested belt in sub-Saharan Africa (Vreysen, 2006).
HAT is caused by two subspecies of Trypanosoma brucei; T. b. gambiense and T. b.
rhodesiense. T. b. rhodesiense is responsible for the acute form (sometimes chronic)
(MacLean et al., 2004) of the disease in Eastern and Southern Africa (Rhodesian HAT),
whereas T. b. gambiense causes the chronic form of the disease in Western and Central
Africa (Gambian HAT) (MacLean et al., 2004; Brun et al., 2010). There is however, a
potential convergence of T. b. gambiense and T. b. rhodesiense foci due to the continued
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expansion of T. b. rhodesiense foci in Uganda towards the northwest (Picozzi et al., 2005)
as farmers move with infected livestock. HAT progresses in two distinct clinical
manifestations, an early stage known as haemolymphatic stage, in which parasites develop
in the tissues, blood and lymph, and the late/ meningoencephalitic stage which occurs
when trypanosomes cross the blood brain barrier and spread to the central nervous system
(Odiit et al., 1997; Kennedy, 2004). Nagana, on the other hand, is caused mainly by
Trypanosoma congolense, Trypanosoma vivax and Trypanosoma brucei brucei. Other
species such as T. evansi, T. godfreyi, T. simiae and T. equiperdum also cause AAT. Mixed
infections involving two or three species have also been reported (Taylor and Authie,
2004). Nagana develops in three stages; anaemia, tissue lesions and immunosuppression.
In the absence of treatment, HAT and AAT are fatal irrespective of the causative species
(Pentreath, 1995).
Control strategies against AT are mainly chemotherapy to treat individuals and animals,
and vector based strategies to block parasite transmission by the fly. Vector control
strategies include use of insecticides (the sequential aerosol technique), bait technologies
(traps and targets), push and pull technique (Hassanali et al., 2008) and the sterile insect
technique (SIT) (Vreysen et al., 2000). Drugs in routine clinical use include pentamidine
and suramin for treating early stage disease and eflornithine and melarsoprol for late stage
HAT disease (Bacchi, 2009). Treatment is dependent on accurate diagnosis and staging of
the disease followed by administration of appropriate drugs. Escalating costs and
limitations in maintenance of tsetse control strategies, however, has hindered vector based
control strategies, and resulted in the reliance of trypanocidal drugs to treat (HAT and
AAT) and/or prevent the disease (AAT). The drugs are however old and faced with
problems of toxicity and ineffectiveness due to resistance (Barrett et al., 2007).
Presently, methods used for sleeping sickness diagnosis include examination of clinical
presentations, parasitological demonstration of the parasites in body fluids, use of
molecular probes for trypanosome subspecies specific nucleic acids (Masiga et al., 1992)
and serological assays targeting trypanosome specific antibodies (Magnus et al., 1978;
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Lejon et al., 1998a; Lejon et al. 1998b) or antigens (Nantulya, 1997). Clinical signs such
as fever, headache, joint pains, and swollen lymph nodes are non specific for HAT,
necessitating the need for laboratory diagnosis. Microscopic demonstration of the parasite
in body fluids is the method recommended by the WHO as the golden standard to diagnose
African trypanosomiasis (Chappuis et al., 2005). Although this method provides direct
evidence for trypanosome presence, it is labour intensive and suffers from low sensitivity
leading to approximately 30% of HAT-positive persons being missed (Chappuis et al.,
2005). Molecular diagnostic tools have high sensitivity and specificity, but require well
equipped health facilities, constant power supply and qualified personnel, resources which
are lacking in most remote rural foci where the disease is endemic (Trouiller et al., 2002;
Brun et al., 2010). Serological tests, such as the Card Agglutination Test for Trypanosomes
(CATT) that involves detection of T. b. gambiense LiTat 1.3 variable antigenic types
(VAT) (Magnus et al., 1978), require minimal equipment and are easily applicable in field
conditions with limited health facilities. A similar kit is however absent for T. b.
rhodesiense diagnosis (Radwanska, 2010). Moreover, clinical signs are insufficient for
specific disease detection, as the signs are nonspecific and mimic other diseases such as
malaria, enteric fever and tuberculous meningitis (Chappuis et al., 2005) that co-exist in
localities. The high toxicity of drugs for late stage disease, the acute nature of Rhodesian
HAT and the limitations with current diagnostic tools necessitate the need for
improvement and/or development of novel diagnostic tools which are simple, specific and
field applicable. This study validated trypanosome surface coat, the variable surface
glycoprotein (VSG), as potential sero-diagnostic candidates for Rhodesian HAT.
African trypanosomes are exclusively extracellular parasites covered by a dense surface
coat monolayer, the VSG (Marcello & Barry, 2007), which shields invariant surface
antigens from immune detection (Cross, 1990a, 1990b; Ferrante & Allison, 1983). The
VSG is highly immunogenic, and forms a potential sero-diagnostic candidate for T. b.
rhodesiense infection. Trypanosomes undergo antigenic variation, a mechanism which
involves change of the VSG to an antigenically different one (Borst & Ulbert, 2001; Borst,
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2002). The parasite’s genome has 1000 - 2000 alternative and highly diverged surface
antigen genes and pseudogenes, of which only a single one is expressed in any individual
trypanosome cell (Lythgoe et al., 2007). Past studies (Barry and Turner, 1991; Frank,
1999; Turner, 1999) have shown that despite the high diversity of antigenic variants,
parasitaemia develop as a series of outbreaks, each outbreak dominated by relatively few
antigenic types. The expression of dominant antigenic variants is not entirely random but
rather sequential and hierarchical (Frank, 1999). The VSG ability to elicit an antibody
mediated immune response, and their sequential expression from a possible >1000 genes,
forms a potential diagnostic target for Rhodesian HAT.
In this study, two variant antigen types (VAT) previously shown to be predominantly
expressed in early phase of T. b. rhodesiense in experimentally infected vervet monkeys
(Cercopithecus aethiops) (Thuita et al., 2008; Masiga et al., unpublished data) were
expressed in bacterial and insect expression systems and purified by immobilized metal
affinity chromatography. Subsequently, the diagnostic potential of the recombinant
antigens were assayed by Ouchterlony double diffusion test against 12 monkey serum
infected with different T. b. rhodesiense strains collected at different days post infection
(dpi) and two uninfected control sera samples. Both recombinant antigens generated from
bacterial and insect expression systems were serologically detected by anti VSG specific
antibodies in the assayed monkey sera indicating their potential use for sero-diagnosis of
Rhodesian HAT, the acute form that kills in weeks or months if untreated.
1.2 Statement of the problem
Current tools used in HAT diagnosis suffer from low sensitivity and specificity, high costs,
and are not applicable in the field environment. Furthermore, the T. b. rhodesiense causes
the acute form of disease, drugs used to treat late stage disease are toxic and currently
there exists no easy-to-use test for this disease. These reasons necessitate early and
accurate diagnosis of the disease to evade complicated and risky treatment procedures.
This need is especially critical as it is anticipated that the number of sleeping sickness
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cases will fall in the next few years, making less sensitive methods unsuitable for tracking
patients during the elimination phase of sleeping sickness. Therefore, there is need for
identification of new diagnostic molecules for diagnosis and staging of the disease.
1.3 Justification
For improved Rhodesian HAT control, there is an urgent need for the enhancement of
current and/or development of new diagnostic tools because of the following reasons.
First, the clinical, parasitological and molecular methods of diagnosis relied on for T. b.
rhodesiense have limitations. Clinical signs are non specific for HAT, parasitological
approaches are low in sensitivity and specificity, while molecular approaches in which
detection of parasite specific nucleic material is targeted, specifically serum resistance-
associated (SRA) gene are not only difficult to apply in screening, but also expensive.
Further, the required resources are not readily available in remote foci where the disease
is prevalent. Secondly, being the acute form with severe clinical symptoms that results
into death within weeks or months, a reliable, sensitive and ready-to-use point-of-care
diagnostic tool(s) is needed for early diagnosis. Otherwise, detection at the late-stage
where the only alternative treatment option, melarsoprol, is inevitable; treatment failures
have been reported and results in 10% of patients developing post-treatment reactive
encephalopathy (PTRE) due to the drug, half of whom die. Thirdly, the potential
convergence of T. b. gambiense and T. b. rhodesiense foci in Uganda due to expansion
necessitates the development of simple and sensitive detection kits specifically for T. b.
rhodesiense. This will be essential in easy diagnosis of mixed infections and
epidemiological surveys. Finally, large scale screening of animal reservoirs of T. b.
rhodesiense is not possible using the current methods. This greatly limits surveillance and
the ability to carry epidemiological studies that are required especially after the recently
reported cases of Rhodesian HAT in Maasai Mara game Reserve in Kenya (Clerinx et al.,
2012; Wolf et al., 2012), a previously unknown focus. Therefore, a VAT-antibody
detection approach could allow development of a robust, simple, ready-to-use and large
scale screening diagnostic kit.
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1.4 Research Hypothesis
Variable surface glycoproteins expressed predominantly during early phase of T. b.
rhodesiense infection do not elicit strong immunological responses and cannot be used as
targets for diagnosis.
1.5 Objectives
1.5.1 General Objective
To evaluate the diagnostic potential of variable surface glycoproteins identified
predominantly early during trypanosomiasis onset as candidates for T. b. rhodesiense
diagnostic tool.
1.5.2 Specific Objectives
i. To generate recombinant variable surface glycoproteins predominantly expressed
during early stages of T. b. rhodesiense infection in bacterial and insect expression
systems
ii. To evaluate the diagnostic potential of the generated recombinant VSG antigens
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CHAPTER TWO
LITERATURE REVIEW
2.1 African Trypanosomiasis
African trypanosomiasis is a disease that affects both humans and animals and is caused
by uniflagellated single-cell extracellular parasites called African trypanosomes (genus,
Trypanosoma). The parasite is transmitted between successive mammalian hosts by the
bite of an infected tsetse fly (genus, Glossina), an insect vector found in 37 countries in
sub Saharan Africa where the disease is endemic. In man, the disease is called sleeping
sickness (or human African trypanosomiasis, HAT) and is caused by two subspecies of T.
brucei, namely T. b. rhodesiense and T. b. gambiense. On the other hand, the disease is
called Nagana or African animal trypanosomiasis (AAT) in domestic animals and is
mainly caused by T. congolense, T. vivax and T. brucei subspp brucei, although other
species including T. evansi, T. equiperdum, T. simiae and T. godfreyi cause AAT.
Trypanosoma vivax and T. evansi also cause disease outside Africa, where they are
transmitted mechanically by biting insects such as tabanids and stomoxes (Osório et al.,
2008; Desquesnes et al., 2013).
2.1.1 African Animal trypanosomiasis (AAT)
African animal trypanosomiasis (AAT) is a parasitic disease caused by African
trypanosomes. This disease affects domestic and wild animals, and is transmitted by tsetse
flies (genus Glossina). An estimated 60 million cattle are at risk of infection in sub-
Saharan Africa where the biological vector exists (Cecchi & Mattioli, 2009). This disease
is endemic throughout the tsetse fly infested regions of Africa (the tsetse belt) covering
over 10 million square kilometres in 37 sub-Saharan African countries (Figure 2-1).
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Figure 2-1. Tsetse fly and cattle distribution in Africa. Areas in red represent the tsetse
infested regions where livestock keeping is constrained, the while the approximate cattle
distribution is illustrated by black dotted areas. Image adopted from International Atomic
Energy Agency (IAEA) (http://allafrica.com/view/group/main/main/id/00012920.html
[accessed 18 Aug 2015]).
Nagana severely affects the development of agriculture in sub-Saharan Africa,
contributing to heavy direct losses due to animal mortality, morbidity, reduced meat and
milk productivity, and reduced calving rates and manure for fertilizing land for crop
production (Vreysen 2006; Cecchi et al., 2014). Nagana also prevents the integration of
crop farming and livestock keeping which is crucial to the development of sustainable
agricultural systems. Notably, this disease limits the exploitation of animal draught power
and transport, and constrains the optimal utilization of fertile lands, reducing food security
(Cecchi et al., 2014). An estimated 3 million cattle deaths are reported annually as reported
by Vreysen (2006). The losses incurred as a result of reduced meat and milk production,
animal death, loss of draught power and costs of programs that attempt to control nagana
are estimated to amount to between 0.6 and 1.2 billion US dollars each year (Budd, 1999).
The overall impact on agriculture Gross Domestic Product loss due to AAT is
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approximated to be 4.75 billion US dollars annually (Budd, 1999; FAO, 2007).
The major pathogenic tsetse-transmitted trypanosome species are T. congolense, T. vivax,
and T. brucei subsp. brucei which affect cattle, goats and sheep, and T. simiae which affect
pigs. Trypanosoma congolense (subgenus Nannomonas) is the single most important
cause of AAT in cattle (Maré, 2004), and affects sheep, goats, horses, and pigs. T.
congolense is grouped into three types namely savannah, Kenya coast/kilifi and
forest/riverine types, which are morphologically indistinguishable. Trypanosoma vivax
(Subgenus Duttonella) also affects cattle, sheep and goats but is less pathogenic for cattle
than T. congolense (Maré, 2004). T. brucei brucei (subgenus Trypanozoon) causes disease
in cattle, sheep and goats. It is morphologically, biochemically and antigenically
indistinguishable to T. b. rhodesiense and T. b. gambiense, but differs in host specificity;
T. b. rhodesiense and T. b. gambiense affects human due to their ability to neutralize
human trypanolytic serum factors TLF1 and TLF2, a characteristic lacking in the other
trypanosome species (Raper et al., 1996; Pays et al., 2001). Trypanosoma evansi is
mechanically transmitted by biting flies such as Tabanidae species and Stomoxys, and in
Africa it mostly affects camels (Desquesnes et al., 2013), causing a disease called
‘‘Surra’’, with cattle being reported as the next highly susceptible host to T. evansi
(Mahmoud & Gray, 1980). Trypanosoma equiperdum affects horses and other equids,
causing a disease termed as “Dourine” (Brun et al., 1998; Claes et al., 2005), which is
transmitted during breeding. Other species such as T. simiae, which infects pigs, and T.
godfreyi, a recently identified species which is pathogenic to pigs (Adams et al., 2010)
also cause AAT. Trypanosoma uniforme and T. suis are other less common tsetse
transmitted species.
Nagana presents itself as an acute, sub acute or chronic disease (Maré, 2004) characterized
by intermittent fever, anaemia, occasional diarrhoea, oedema of limbs and genitalia, and
often terminates in death in the absence of treatment. The acute form of the disease causes
abortion, central nervous system disorders and death, while in the chronic condition which
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is the most common form of the disease and often aparasistaemic, affects the working
capacity and productivity of the animals, causes severe anaemia, weight loss and infertility
(Taylor & Authie, 2004).
Trypanocidal drugs available for AAT include isometamidium chloride (trade names:
Samorin, Trypamidium, MandB 4180A) which is used as a prophylactic drug, while
diamazene aceturate (trade name: Berenil) is used for treatment. These drugs are over a
century old, and they are ineffective due to increasing resistance (Chitanga et al., 2011),
necessitating the need to identify novel compounds for treatment of the disease.
Clinical signs are non specific for AAT diagnosis and cannot be relied upon solely,
requiring the need for laboratory confirmation. Demonstration of trypanosomes in body
fluids is the standard diagnostic technique used, a technique which suffers from low
sensitivity and specificity leading to half of the infected animals being missed (Nantulya,
1990). Sample inoculation in rats or mice has been used to diagnose AAT, a method that
enables detection of low levels of parasites, but this method is time consuming, costly and
non amenable to such species which do not infect laboratory rodents such as T. vivax, T.
simiae and T. congolense (Nantulya, 1990). Detection of anti-trypanosome antibodies in
blood samples using indirect fluorescence antibody test (IFAT) and enzyme linked
immonosorbent assay (ELISA) are alternative diagnostic tests used, but suffer from low
specificity because of cross reactivity with non-target parasitic diseases, require
sophisticated laboratory equipment thereby limiting their application in the field
(Nantulya, 1990). Detection of trypanosome antigens in circulation rather than antibodies
elicited after infection enables the differentiation of current infection from successfully
treated or self cure cases. Antigen detection tests using polyclonal antibodies against T.
vivax and T. congolense antigens (Rae & Luckins, 1984) and monoclonal antibodies
(Nantulya et al., 1987) have been applied in the diagnosis of AAT using IFAT and
sandwich ELISA tests. Polyclonal antibodies have however been shown to be limited by
cross reactivity with non target trypanosome species and other blood parasitic animal
11
diseases leading to low specificities, while species specific monoclonal antibodies
recorded high sensitivity (92%) and specificity (100%) with no cross reactions between
trypanosome species (Nantulya et al. 1987; Nantulya 1990). Antigen detection tests are
more sensitive than the recommended parasitological detection tests and can be applied in
large scale screening, but suffer from their inability to detect infections at onset of disease
before the parasites are destroyed to release the antigens for detection (Nantulya &
Lindqvist, 1989).
2.1.2 Human African Trypanosomiasis (HAT)
Human African trypanosomiasis (HAT) or sleeping sickness is a disease caused by two
subspecies of Trypanosoma brucei, T. b. gambiense and T. b. rhodesiense. Trypanosoma
b. gambiense is responsible for the chronic form of the disease in Central and Western
Africa (Gambian HAT) while T. b. rhodesiense causes an acute form of the disease in
Eastern and Central Africa (Rhodesian HAT) (figure 2-2). There is, however, a risk of
overlap of the two forms of disease in Uganda due to expansion of T. b. rhodesiense foci
(Picozzi et al., 2005), with major impacts on diagnosis and treatment of the disease (Brun
et al., 2010).
In the tsetse fly belt, about 70 million people are at risk of infection, and an estimated
3,796 new cases were reported in 2014 (WHO, 2015) compared to 7,214 new cases
reported in 2012 (WHO, 2013). This represents a 90% reduction in cases reported as
compared to the peak which occurred in 1998 when 37,991 new cases were reported
(Simarro et al., 2012). This could however be an underestimate as most cases occur in
remote rural areas mostly in Democratic Republic of Congo, the Republic of Congo,
Angola, Central African Republic and Southern Sudan (WHO, 2001; Brun et al., 2010)
where health infrastructures are poor, leading to most cases being missed due to lack of
appropriate diagnostic tools, or unreported due to inaccessibility of some areas due to
geopolitical unrests e.g. in Southern Sudan.
12
The clinical manifestation of HAT involves two stages. The first stage of the disease (the
haemolymphatic stage) characterized by presence of the parasite in blood and lymph fluids
is accompanied by fever, headache, adenopathy, joint pain and pruritus. Rapid parasite
growth is countered by host immune responses, but parasite antigenic variation, a
mechanism that involves the parasites continual switching from the expression of one
variant surface glycoprotein (VSG) on the surface to the expression of a different
immunologically distinct VSG (Donelson, 2003), enabling the parasites’ persistence in the
hosts blood stream.
Subsequently, the parasites cross the blood-brain barrier into the cerebrospinal fluid
causing the second stage (encephalitic stage) of the disease (de Atouguia and Kennedy,
2000). At this stage, sleeping sickness causes an alteration of the circadian sleep/wake
cycle, and is accompanied by endocrinological, cardiovascular and renal disorders, which,
in the absence of treatment, progress to body wasting, somnolence (hypersomnia), coma
and ultimately death of the patient (Kennedy, 2006, 2013).
13
Figure 2-2. Human African trypanosomiasis (HAT) distribution, incidence and risk
for travelers in sub Saharan Africa. The black line divides the Gambian and Rhodesian
disease foci within the geographic distribution of tsetse fly. Image adopted from Brun et
al. (2010).
2.2 Trypanosome life cycle
African trypanosomes are digenetic, alternating between vertebrate mammalian and
invertebrate tsetse fly hosts (Figure 2-3). During a blood meal, an infected fly injects
metacyclic trypomastigotes into blood and subsequently the lymphatic system of
mammalian host. In the bloodstream, the metacyclic trypomastigotes transform into
bloodstream trypomastigotes (morphologically slender forms) and are carried to other
sites throughout the body where they continue to multiply. The blood stream
14
trypomastigotes express the bloodstream-stage-specific VSG coat to evade the
mammalian immune response via antigenic variation (Matthews, 2005). At high densities,
differentiation to morphologically stumpy forms occurs; these are division-arrested forms
pre-adapted for transmission to tsetse flies (Matthews, 2005; Denninger et al., 2007).
Upon uptake during feeding, the stumpy forms transform into procyclic forms, which are
the proliferative forms in the fly’s midgut, and express a surface coat called procyclin that
is distinct from the bloodstream VSG coat. After establishment in the fly midgut, division
is arrested and the trypanosomes migrate to the tsetse salivary gland, where they attach as
epimastigote forms. These are proliferative and attached through elaboration of their
flagellum. Eventually, the epimastigotes generate non-proliferative metacyclic forms,
which re-acquire a VSG coat in preparation for transmission into a new mammalian host
(Matthews, 2005). Theparasites cycle in the fly takes about 2 to 3 weeks (Chappuis et al.,
2005).
15
Figure 2-3. Life cycle of Trypanosoma brucei in mammalian host and the tsetse fly
vector. The different developmental stages of trypanosomes in the mammalian blood
stream (blood stream slender and stumpy forms), tsetse flies midgut (procyclic) and
salivary glands (epimastigote and metacyclic stages, respectively) is illustrated, showing
the surface coat at each stage. Image adopted from Matthews (2005).
2.3 Control of African trypanosomiasis
Various methods have been used to control trypanosomiasis, with such methods as vector
based control strategies, screening of populations and curative treatment of infected
people being used for HAT management. Management of AAT involves prophylactic and
curative treatment of animals with trypanocidal drugs, the promotion of trypanotolerant
cattle and suppression or eradication of the vector (Vreysen et al., 2012).
16
2.3.1 Chemotherapy
Chemotherapy for HAT has lagged behind, with the current drugs in use being more than
50 years in existence (Chappuis et al., 2005; Bacchi, 2009). Treatment of sleeping sickness
is complicated and depends on the T. brucei subspecies involved and on the stage of the
disease (Radwanska, 2010).
2.3.1.1 Drugs for Early Stage Treatment
Pentamidine is a water soluble aromatic diamine (diamidine) developed for treatment of
early stage T. b. gambiense, leishmaniasis and pneumonia (Nok, 2003). A dosage consists
of 7-10 intramuscular injections (Bacchi, 2009). This drug however causes severe side
effects such as hypoglycaemia, hyperglycaemia and hypotension (Kennedy, 2006b).
Suramin is a sulphonated naphthylamine agent for treatment of early stage T. b.
rhodesiense disease. The inability of suramin to penetrate the blood brain barrier renders
this drug ineffective in the treatment of second stage disease. Suramin treatment regimen
includes five intravenous injections every three to seven days for four weeks (Etet &
Mahomoodally, 2012).
2.3.1.2 Drugs for Late Stage Treatment
Melarsoprol, an intravenously administered drug, is an arsenic drug used in first line
therapy for late stage Gambian and Rhodesian HAT. Earlier, the standard melarsoprol
regimen included two to four courses of intravenous injections, each course consisting of
three injections administered after a week’s break between the courses. Recently, a ten
day continuous regimen was introduced for treatment of Gambian HAT, and has been
adopted in many endemic countries (Kennedy, 2006b). This drug is painful to administer,
destroys veins after several applications and is faced with toxicity caused by reactive
arsenical induced encephalopathy in 10% of patients it is administered to, followed by
pulmonary oedema and death in more than half of the cases in 48 hrs (Chappuis, 2007;
Bacchi, 2009). Treatment failure rates have also been reported in DRC, CAF, Sudan and
Uganda (Brun et al., 2001). Melarsoprol is also administered for treatment of late stage T.
b. rhodesiense disease. Eflornithine (α-difluoromethylornithine or DFMO) is a recently
17
developed alternative drug for treatment of late stage T. b. gambiense disease. Eflornithine
drug regimen is long, and doses are given by prolonged intravenous infusion (Etet &
Mahomoodally, 2012).
2.3.1.3 New Drugs in Use or Under Clinical Trials
New drugs are currently under preclinical and/or clinical trials, while a new combination
therapy has been introduced for treatment of second stage Gambian HAT. A Diamine
derivative (CPD-0802) entered advanced preclinical trials in mouse models for second
stage HAT treatment, Nitroheterocycle derivative (fexinidazole) entered phase II clinical
trials, and a Benzoxaborole derivative (SCYX 7158) entered phase I clinical trials (Barrett
& Croft, 2012). These compounds are still under considerations for further development
(Thuita et al., 2015).
The introduction of Nifurtimox-Eflornithine Combination Therapy (NECT) in 2009 for
treatment of second stage T. b. gambiense HAT has been shown to be less toxic and more
effective compared to eflornithine only treatment regimen in a preclinical study conducted
in Congo (Priotto et al., 2007).
2.3.2 Vector Control
Trypanosome transmission blocking by vector based control strategies is the most
preferred method of controlling the disease due to increased tolerance to trypanocidal
drugs in use. The adult fly is mostly targeted as it is the only accessible stage of the fly
due to the absence of eggs, a larval free stage in nature and the fact that the pupa develops
in the soil (Vreysen et al., 2012). Removal of the vectors’ preferred vegetation and the
killing of host game to control tsetse flies proved to be a very efficient vector control
strategy, but this strategy has become unacceptable due to its negative effect on the
ecosystems.
Pour flow strategy of residual insecticides such as dichlorodiphenyltrichloroethane
(DDT), dieldrin and endosulfan in areas where tsetse flies rest during dry seasons was
18
effective in controlling the disease, but has been limited due to logistical difficulties
(supervision and planning) and labour intensiveness. The use of insecticides which often
persist for long periods of time on the environment after spraying leads to the potential
development of resistance to the insecticide, killing of non target insects, the potential
outbreak of other pests due to elimination of their natural predators and the destruction of
beneficial insects such as bees, ladybugs and beetles involved in pollination. The
cumulative effect of insecticides also increases general pollution of the environment
because of accumulation of the chemicals in the insecticides (Vreysen et al., 2012).
The application of persistent insecticides has been gradually replaced by the aerial
spraying of non residual ultra-low volume insecticides with planes, a technique known as
the sequential aerosol technique (SAT) that is more environmentally friendly. The
insecticide drop-size is small enough to suspend in the air but heavy enough to prevent
upward drift (Vreysen et al., 2012). Electrically or air driven rotary atomizers are used at
a speed of 16,000 rpm to produce an aerosol of droplets of 30–40 µm. This control tactic
aims at killing all adult flies in each spraying cycle through direct contact with the
insecticide mist followed by all emerging flies in the subsequent cycles before they can
start reproducing.
A technique that targets tsetse flies using stationary attractive devices such as cloth
traps/targets that either kill the flies through tarsal contact with insecticides applied to the
surface of the target or by heat or starvation after being guided to a non-return cage
(Vreysen et al., 2012) has been applied as a control strategy. The trap method is relatively
cheap, unsophisticated and can be improved by incorporating insecticides such as
pyrethroids. A live bait technique which involves insecticide treatment of livestock
exploits the blood sucking behaviour of both sexes of tsetse which are killed by picking
up a lethal deposit of insecticide on the ventral tarsal spines and on pre-tarsi during
feeding. This method requires no sophisticated equipment, and insecticide application is
rapid and easy (Vreysen et al., 2012).
19
Sterile insect technique (SIT) involves the mass rearing of a target insect, sterilization by
exposure to gamma or X rays or chemicals and the subsequent and sustained release of
the sexually sterile insects into the target population (Dyck et al., 2005) to outcompete the
indigenous insect for the female insects resulting in no production of offsprings (Vreysen
et al., 2013). This technique was applied in Zanzibar (Ugunja island) in the eradication of
Glossina austeni using gamma sterilized male flies from 1994 to 1997 (Vreysen et al.,
2000). The SIT is environment friendly, has no adverse effects on non-target organisms,
is species-specific and can easily be integrated with biological control methods such as
parasitoids, predators and pathogens. There is no evidence of development of resistance
to the effects of the sterile males provided that adequate quality assurance is practiced in
the production process. This technique is, however, only effective when the target
population density is low, requires detailed knowledge of the biology and ecology of the
target insect, and the insect should be amenable to mass-rearing (Vreysen et al., 2013).
Combinations of repellent and attractant chemicals have also been used in push-pull
tactics. Tsetse flies show a feeding preference on different vertebrate animals and appear
to use push–pull signals actively to avoid some hosts and to locate those preferred. This
strategy involves the use of synthetic repellents to push the flies away, and to attract them
(pull) into baited traps (impregnated with insecticides). Natural and synthetic repellents
such as 2-methoxyphenol, a constituent of bovid odors offer less protection to cattle
compared to a repellent obtained from waterbuck, Kobus defassa, which is refractory to
tsetse flies (Hassanali et al., 2008).
One novel control strategy under development involves reducing the ability of the tsetse
fly to transmit trypanosomes by expressing anti-trypanosomal molecules in Sodalis
(Sodalis glossinidius), the commensal symbiont in the tsetse midgut. This is an
environmentally acceptable, efficacious and affordable technique for the control of tsetse
flies (Medlock et al., 2013).
In sum, vector control strategies targeting the adult stage of the fly have recorded mixed
success in controlling African trypanosomiasis, complementing the use of trypanosomal
20
drugs which have been limited by increasing tolerance and unacceptable toxicity.
Vegetation clearance and insecticides have an adverse effect on the environment, and the
application of SIT, live bait technique, push-pull technique and paratransgenesis offer a
more environmentally friendly options. However, the application of vector control
strategies are limited by logistical hurdles. Therefore, there is need for improvement of
vector control methods, development of new safe drugs and novel diagnostic tests for
African trypanosomiasis.
2.4. Diagnosis of Human African Trypanosomiasis
Despite its huge impact, human African trypanosomiasis is a neglected tropical disease,
with little progresses in development of novel drugs and diagnostic tools (Papadopoulos
et al., 2004). Diagnosis is limited by low sensitivities and specificities of the current
diagnosis procedures (Radwanska, 2010). The high toxicity of melarsoprol for stage 2
treatment necessitates high accuracy in both diagnosis and staging of the disease
(Chappuis et al., 2005). The approaches used in the diagnosis of HAT are discussed below.
2.4.1. Clinical Diagnosis
Clinical diagnosis entails the examination of individuals for clinical signs such as enlarged
cervical lymph nodes (lymphadenopathy), excessive collection of fluids in tissues
(oedema), enlarged spleen and liver (splenomegaly), fever, pruritus, development of a
chancre at the site of tsetse bite and joint pains (Chappuis et al., 2005; Radwanska, 2010).
Early stage symptoms are experienced 1-3 weeks after the bite of an infected tsetse fly,
but these clinical signs are in general insufficient for specific diagnosis of sleeping
sickness, leaving most patients undiagnosed at the early stage of disease or misdiagnosed
for other endemic and more frequent tropical diseases such as malaria (Kennedy, 2013).
Owing to these varied clinical manifestations, diagnosis of trypanosomiasis cannot be
solely based on clinical signs alone to both confirm or to stage the disease. Limitations in
clinical diagnosis necessitate laboratory confirmation (Nantulya, 1990) and staging of
21
HAT disease using serological tests targeting anti trypanosome specific antibodies raised
by patients’ immune response to screen populations at risk, microscopy to demonstrate
the presence of trypanosomes in body fluids and/or molecular amplification to detect
parasite specific nucleic acids.
2.4.2 Parasitological Diagnosis
Parasitological diagnosis involves the microscopic detection of trypanosomes in blood, in
chancre aspirates or cervical lymph node (CLN) aspirates, and cerebrospinal fluid for
stage determination. Trypanosomes in the chancre are detectable days earlier than in blood
by microscopy as fresh, fixed or Giemsa- stained preparations, but because the chancre
disappears before most patients are tested, it is rarely used (Chappuis et al., 2005). The
gold standard for diagnosis of HAT recommended by the WHO is the demonstration of
the presence of parasites in body fluids, a method that suffers from low sensitivity and is
cumbersome. About 20 – 30% of patients are estimated to be missed by the various
parasitological tests in use (Robays et al., 2004; Chappuis et al., 2005). The standard
parasitological techniques applied in the diagnosis of HAT are discussed below.
2.4.2.1 Wet Blood Films
This technique is used to examine fresh blood between a cover slip and a slide with a
microscope. The parasites are seen either directly moving between the blood cells or
indirectly as they cause the blood cells to move. This technique is simple and inexpensive,
offering immediate diagnosis upon parasite detection. However, this technique has limited
sensitivity with a detection limit as high as 10,000 trypanosomes per ml (corresponding
to 1 parasite in 200 microscope fields) (Chappuis et al., 2005). This technique requires
fresh blood samples as the trypanosomes lose mobility after time, and cannot be used to
stage the disease nor identify the trypanosome species (Chappuis et al., 2005). The blood
films can either be thick or thin blood films.
22
2.4.2.2 Microhematocrit Centrifugation Technique (mHCT)
This technique, also referred to as capillary tube centrifugation (CTC) technique or the
Woo test, involves the use of capillary tubes containing anticoagulant which are filled with
finger prick blood, one end sealed with plasticine and the trypanosomes concentrated by
high speed centrifugation in a hematocrit centrifuge for 6-8 minutes. The parasites are
concentrated at the level of leukocytes, between the plasma and the erythrocytes (Woo,
1969; Chappuis et al., 2005), and the capillary tubes can be directly examined at low
magnification of ×100 or ×200 for mobile parasites. This test has a detection threshold of
500 trypanosomes per ml. In addition to being time consuming, the presence of
microfilaria (the pre-larval stage of Filarioidea in the blood of humans) limits
visualization of trypanosomes.
2.4.2.3 Quantitative Buffy Coat (QBC) technique
Originally developed for the assessment of differential cell count, this technique has been
used to concentrate and stain the nucleus and kinetoplast of trypanosomes with acridine
orange dye, clearly discriminating the parasites from leukocytes. Blood is centrifuged at
high speed in capillary tubes with ethylenediaminetetraacetic acid (EDTA) and acridine
orange containing dye, enabling the discrimination of the parasites from leukocytes.
Motile trypanosomes can be identified by their fluorescent kinetoplasts and nuclei in the
expanded buffy coat using ultra violet light in a dark room. This technique has a high
sensitivity, with 95% sensitivity for parasite concentrations of 450 parasites per ml, and
can detect more patients with low parasitaemia than the mHCT (Chappuis et al., 2005).
However, the fragility and sophistication of the materials used in this technique limits its
application in the field, and thus is not used most frequently in field diagnosis.
2.4.2.4 Mini-Anion-Exchange Centrifugation Technique (mAECT)
This technique is based on use of anion exchange chromatography to separate
trypanosomes from blood cells since the parasites are less negatively charged than blood
23
cells. The blood is first passed through anion exchange gel column, separating the
parasites from the blood, with parasites collected in a sealed glass tube. The parasites are
subsequently concentrated at the bottom of the sealed glass tube by low speed
centrifugation. The tip of the glass tube is then examined in a special holder under the
microscope for presence of parasites. The large blood volume used (300 µl) enables the
detection of 100 trypanosomes per ml resulting in high sensitivity, but this technique
involves tedious manipulations and is time consuming.
2.4.3 Serological Diagnosis
Tests for screening the population at risk in T. b. gambiense prone foci are mostly based
on antibody detection against the trypanosomes’ VSGs, although tests that detect
trypanosome antigen against trypanosome specific antibodies have also been developed.
Serological tests require minimum equipment and are easily applicable in the field
environment with limited health facilities. Serological test used in the diagnosis of HAT
are discussed below.
2.4.3.1 Immunofluorescence Antibody Test (IFAT)
The immunofluorescence antibody test (IFAT) is a serological test for both serum and
dried whole blood on filter papers which utilizes IgG antibodies conjugated to fluorescein
isothiocyanate. Before the introduction of CATT, the IFAT was the widely applied
serological test in the mass screening of T. b. gambiense infection in the 1970s (WHO,
2001).
The test procedure includes antigen preparation by inoculating T. b. gambiense in guinea
pig or albino rats, blood collected after the parasitaemia reaches 2-20 parasites per field;
and mixed with heparine-glucose mixture and used to prepare thin blood films consisting
of 5-50 parasites/field. This is followed by placing of patient serum on a glass plate with
the prepared antigen, the mixture is washed with phosphate buffered saline (PBS) and
fluorescein labeled antihuman at a predetermined dilution added. After washing as before,
24
the slides are covered with glycerine-PBS mixture and the slides viewed under UV
microscopes (Wery et al., 1970). IFAT is easily performed by non specialized laboratory
technicians with a read out time of 10 to 30 seconds. However, this test is costly and only
amenable to some laboratory settings.
2.4.3.2 CATT/T. b. gambiense and its variants
The card agglutination test for trypanosomiasis (CATT) is a serological test for diagnosis
of Gambian HAT developed in 1978 by Magnus and colleges (Magnus et al., 1978). CATT
is the most widely used test applied for both passive and active screening of populations
in T. b. gambiense endemic foci due to its simplicity and cost effectiveness. This test is
based on a single lyophilized laboratory adapted strain of T. b. gambiense expressing
predominant variable antigenic type (VAT) designated LiTat 1.3 on the parasites surface.
Applied on whole blood, plasma or serum, the CATT/ T. b. gambiense is used in active
screening of populations at risk, and seropositive cases are further confirmed by
parasitological examination.
Mass production of trypanosomes expressing the variable antigenic type LiTat 1.3 antigen
is achieved by culturing the parasites in laboratory rodents and the blood stream stage of
the parasite extracted from the blood; a process which puts laboratory staff at risk of
infection (Van Nieuwenhove et al., 2012). The harvested parasites expressing VAT LiTat
1.3 are fixed, stained with Coomassie blue before being lyophilized (Chappuis et al.,
2005). The kit consists of the antigen reagent, a control sera and a rotator; and operates by
mixing blood and PBS-resuspended antigen reagent in equal volumes (one drop each), the
mixture is rotated at 60 rpm for 5 minutes using the rotator and the results analyzed by
macroscopic viewing, and the reported sensitivity and specificity of this test is 87-98%
and 95% respectively (Chappuis et al., 2005). CATT on diluted blood is more cost
effective than on whole blood (CATT/wb), but its slight complication makes it rather
difficult to perform under field conditions (Elrayah et al., 2007).
Micro-CATT test which permits utilization of minute amounts of dried blood samples on
25
filter papers or diluted blood was recommended by WHO for mass screening due to its
affordability (Miezan et al., 1991). This test uses micro-volumes, unlike the classical
CATT/T. b. gambiense test (a fifth of the standard quantities of antigen and sample). This
test is however less sensitive than the classical CATT/T. b. gambiense especially when the
FP elute is older than a day or stored at 4°C (Chappuis et al., 2002; Truc et al., 2002).
Coupled to the decrease in sensitivity with time, micro-CATT also suffers from difficulties
in interpreting agglutination test results due to the micro volumes used in this test
(Chappuis et al., 2002; Truc et al., 2002).
CATT on blood impregnated on filter papers (CATT-FP) was developed at the Department
of Parasitology of the Prince Leopold Institute of Tropical Medicine, Antwerp (ITMA), is
an improvement of the micro-CATT, and involves using a higher volume of the FP eluate
(30 µL). CATT-FP has also been improved to enable the screening of populations in areas
outside active screening radius (Chappuis et al., 2002). The maiden test evaluation of this
test was carried out in south-Sudan in Kajo-Keji County, and compared well with CATT
on plasma, showing a sensitivity of 94% on parasitologically confirmed cases. More so,
the CATT-FP test illustrated that samples could be stored at ambient temperatures (25-
34 °C) and under refrigeration (2-10 °C) for upto fourteen days to eight weeks without
loss of sample integrity (Chappuis et al., 2002), unlike micro-CATT test whose sensitivity
decreased with time. This can be attributed to the strict dry storage of the filter papers
impregnated with blood, ensuring integrity of the blood samples.
Another alternative to the classical CATT/T. b. gambiense test, the CATT-EDTA test
(Pansaerts et al., 1998) is an improvement of the classical CATT/T. b. gambiense test; the
EDTA (10mM) incorporated in the reaction buffer helps reduce complement mediated
prozone effect, increasing the sensitivity of CATT. Active complement present in freshly
collected blood inhibits the agglutination reaction leading to the formation of a prozone
common in classical CATT test (Jamonneau et al., 2000). Dilution of blood also helps
increase the specificity of the CATT-EDTA test (Jamonneau et al., 2000). The CATT-
26
EDTA test was evaluated in a preliminary study conducted in Cote d’Ivoire (Jamonneau
et al., 2000), and showed a higher specificity (94.6%) compared to that of CATT/T. b.
gambiense (92.5%).
A limitation of CATT/ T. b. gambiense screening tests is the observed absence of VAT
LiTat 1.3 in some T. b. gambiense foci e.g. Cameroon (Dukes et al., 1992), leading to false
negative results. In a study carried out in Fontem, Cameroon, it was observed that the
LiTat 1.3 gene was not expressed by T. b. gambiense strains, while a LiTat 3 like gene,
expressed by some strains, elicited antibodies which could not be detected by the CATT
test (Dukes et al., 1992). This lowers the sensitivity of the CATT/T. b. gambiense in such
areas where LiTat 1.3 is not expressed.
2.4.3.3 LATEX/T. b. gambiense
The LATEX/T. b. gambiense utilizes the combination of three predominant VATs, namely
LiTat 1.3, LiTat 1.5 and LiTat 1.6 (VAT G16/6) of bloodstream T. b. gambiense parasites.
The incorporation of three VATs predominantly expressed during T. b. gambiense
infection broadens the geographic user-range of this test. Partially purified antigens are
adsorbed on latex particles/beads, and equal amounts of diluted blood samples are mixed
with the latex reagent and spread onto a 1.5 cm reaction zone with a dark background
(Jamonneau et al., 2000). Agitating the card at 70 rpm for five minutes enables the
antigens to agglutinate with the VAT specific antibodies present in the blood, which are
visible as white macroscopic agglutinations (Büscher et al. 1991; Lejon et al. 1998).
The specificity of LATEX/T. b. gambiense test was reported to be 98.1% in a study
conducted in Cote d’Ivoire. Despite being highly specific, this test requires qualified
personnel, is time consuming and the requirement of microplates complicates its
application in the field setting (Jamonneau et al., 2000). In addition, mixing more than
one antigen in the same reaction reagent reduces the reactivity of the individual antigens
in the final mixture, reducing the sensitivity of the test.
27
2.4.3.4 Card Indirect Agglutination Test for Trypanosomiasis (CIATT).
The CIATT is an antigen detection test which enables the differentiation between active
and cured HAT by the detection of antigens released by non-circulating trypanosomes
sequestered in the liver, spleen, lymph nodes, or CNS (Chappuis et al., 2005). Developed
in Kenya, the card indirect agglutination test for trypanosomiasis (CIATT) or TrypTect
CIATT® is a latex agglutination test for both T. b. gambiense and T. b. rhodesiense
infection first described in 1997 by (Nantulya, 1997). This test detects trypanosome
antigens circulating in the bloodstream using a monoclonal antibody specific to an
invariant internal antigen present in both human infective T. brucei subspecies (Nantulya,
1997) - the antibody is coupled to latex particles, forming the reaction reagent. The assay
involves mixing on the test card equal volumes (50 µl) of test reagent and serum/ plasma,
and after two minutes of incubation the test card is tilted and rotated, macroscopically
showing presence of agglutination when the test sample is positive.
Analysis of parasitologically confirmed 244 T. b. gambiense samples from Uganda and
Côte d'Ivoire, and 132 T. b. rhodesiense samples from Uganda by the CIATT illustrated
that the test was highly sensitive, recording 95.8 % and 97.7 % sensitivities for T. b.
gambiense and T. b rhodesiense infections respectively (Nantulya, 1997). The antibody
coupled to latex particles did not agglutinate with 193 negative sera samples assayed,
illustrating the high specificity of this test (100% specificity). This study showed that the
CIATT was more sensitive than some parasitological tests done in parallel (lymph node
aspirates, mAECT, mHCT and CSF single and double centrifugation techniques).
However, this test could not detect minute antigens in circulation, necessitating the need
to test suspect cases after one to two weeks (Nantulya, 1997).
Antigen detection tests enable detection of active infection, unlike antibody detection tests
which can detect persistent antibodies still in circulation months after the infection has
been cleared due to treatment or self-cure. This means that the CIATT can be used in post
treatment follow-up, and the ability to assess post treatment follow up with finger prick
28
blood could eliminate the need for lumber puncture, a painful and intrusive method
undergone by patients during post treatment follow ups.
2.4.3.5 Enzyme-linked immunosorbent assay (ELISA)
The enzyme-linked immunosorbent assay (ELISA) is an immunological test that
indirectly demonstrates the presence of an infecting parasite in body fluids. In contrast to
CATT and LATEX tests, ELISA makes use of markers to enable the
detection/visualization of antigen–antibody complexes. ELISA has been used in the
serodiagnosis of sleeping sickness to detect anti-trypanosome specific antibodies in serum
and CSF (Lejon et al. 1998) or saliva (Lejon et al., 2003). ELISA has also found
application in the staging of the disease, as anti-trypanosome specific antibodies or
cerebrospinal IgM immunoglobulin whose levels are elevated in the CSF during infection
(Greenwood and Whittle, 1973; Whittle et al., 1977; Lejon et al., 2002).
Trypanosome antigens are prepared by infecting laboratory rodents, and the trypanosomes
are collected by differential centrifugation, resuspended in saline, lysed via sonication and
centrifuged to remove insoluble cell debris (WHO, 1976). The test antigens (crude or pure)
are adsorbed on microplates at different dilution factors, and diluted sera/CSF added. An
enzyme conjugate (antigen specific antibody conjugated to an enzyme such as alkaline
phosphatase or horse radish peroxidase) subsequently added followed by the enzymes
substrate. The optical density is read at a specific wavelength depending on the substrate
used, with the extinction value obtained being a measure of the quantity of enzyme-labeled
immunoglobulin bound to the antigen – antibody complex (Knapen, 1977).
ELISA/T. b. gambiense is highly sensitive (Knapen, 1977) and specific (98.5%) (Elrayah
et al., 2007), cost effective and has acceptable reproducibility. However, this test suffers
from variable sensitivities, and can only be applied in reference centers with highly skilled
technicians, who are more often lacking in poor remote areas. In addition, cross reactions
with antibodies from patients infected with Leishmaniasis is a concern, leading to false
positives (Knapen et al., 1977). A sensitive micro-ELISA for T. b. rhodesiense is also
29
described by (Voller et al., 1975), but this test too suffers from cross reaction by antibodies
specific to, and antigens of Leishmania donovani and T. cruzi.
ELISA based on antigens for sleeping sickness diagnosis is sensitive and not limited by
cross reactions with sera collected from patients infected with common tropical diseases.
In addition, detection of trypanosome specific antigens circulating in the body fluids
provides direct evidence of active infection (Komba et al., 1992). Using a monoclonal
antibody elicited against invariant surface protein at the procyclic stage of the parasites,
(Nantulya, 1989) describes an indirect ELISA tool for diagnosis of Rhodesian HAT. In
this study, the antigen ELISA was 100% specific, and 90% sensitive, and could also detect
six samples which were parasitologically negative. However, this test could not detect
some parasitologically confirmed cases, a limitation attributable to low amounts of
antibody produced or during early stages of the disease (Nantulya, 1989).
2.4.3.6 Immune trypanolysis test (TL)
The immune trypanolysis test (TL), is an antibody-compliment mediated serological test
that detects antibodies raised against specific VATs, revealing/demonstrating prior contact
with the parasite. The variable glycoprotein surface coat of the trypanosomes, being highly
immunogenic, elicits an immune response by the host’s immune system which involves
the production of antibodies specific to the VATs. These antibodies are able to opsonize,
agglutinate and lyse circulating trypanosomes (Van Meirvenne et al., 1995). This test can
only be applied in reference laboratories, limiting its field applicability.
Immune trypanolysis test involves the intravenous inoculation of 107 mouse adapted
bloodstream trypanosome clones expressing predominant VATs and a control VAT in
complement rich cavia serum. Human plasma sample (25 µL) is mixed with 25 µL of
complement rich serum, and 50 µL of 107 trypanosomes/mL suspension prepared from
adapted bloodstream form trypanosomes in mice added. The micro-plates are mixed and
incubated at room temperature for one and a half hours, and the reaction mixture examined
under a phase contrast microscope (Jamonneau et al., 2010).
30
In 1990, Isharaza and Van Meirvenne described a trypanolysis test for the detection of T.
b. rhodesiense antibodies in 85 sera samples collected from Uganda (Isharaza & Van
Meirvenne, 1990). The study used a panel of twelve T. b. rhodesiense specific VATs (ETat
1/1, 1/2, 1/3, 1/10, 1/14, 1/18, 1/19, UTat 3/1, 3/7, 1/1 and 4/1) and one control VAT (AnTat
25/1). All the assayed VATs were able to detect all the seropositive sera samples albeit
with varying proportions. In combination, three of the VATs i.e. ETat 1/1, ETat 1/4 and
UTat 1/1 showed positive reactions with all sera samples infected with T. b. rhodesiense.
However, some parasitologically positive sera were missed by the assayed VATs; this can
be attributed to low levels of anti VAT specific antibodies in circulation (Isharaza & Van
Meirvenne, 1990).
Van Meirvenne and colleagues described a study where a panel of twelve predominantly
expressed T. b. gambiense VATs were used (LiTat 1.1 to LiTat 1.10, and AnTat 11.17 and
AnTat 11.20), and a control from T. b. rhodesiense; ETatR1 (Van Meirvenne et al., 1995).
This study was highly specific (100%) in different studies conducted in T. b. gambiense
endemic countries such as Equatorial Guinea, Côte d'Ivoire, Gabon, Congo, Uganda
(north west part of the country), Zaire, Sudan and Nigeria. However, the sensitivities vary
depending on the sera dilution factor (Van Meirvenne et al., 1995). In addition, some of
the VATs included in this study were absent in some countries, narrowing the geographical
user range of the test.
In 2014, Camara et al. evaluated the use of blood spotted filter papers as a substitute for
serum or plasma in immune trypanolysis test. Blood spotted filter papers can be stored
and transported under field conditions, while serum or plasma require a cold chain.
Notably, their results showed that use of filter papers resulted in decline in sensitivity, but
specificity was not affected (Camara et al., 2014).
2.4.4 Molecular Tools
Molecular tools, specifically polymerase chain reaction (PCR), have been applied in the
detection of trypanosome specific nucleic acids (Masiga et al., 1992; Kabiri et al., 1999;
31
Radwanska et al., 2002a; Radwanska et al., 2002b; Desquesnes and Dávila, 2002; Gibson,
2002, 2009; Adams and Hamilton, 2008). Generic primers that allow detection of multiple
trypanosome species by targeting a common internal transcribed spacer (ITS)
(Desquesnes et al., 2001; Njiru et al., 2005) exhibits interspecies size variation. While ITS
amplification is more applicable in screening, it cannot distinguish the members of T.
brucei, a limitation suffered by amplification of kinetoplast minicircle (Mathieu-Daudé et
al., 1994) and mini-exon repeat DNA (Ramos et al., 1996), hence the need for subspecies-
specific amplification.
PCR targeting single copy sequences for the diagnosis of Rhodesian HAT based on the
detection of serum resistance associated (SRA) gene (Gibson et al., 2002; Radwanskaet
al., 2002a) and the diagnosis of Gambian HAT by targeting the Trypanosoma gambiense
specific glycoprotein gene (TgsGP) (Radwanska et al., 2002b) have been used on blood
samples from CATT positive patients, and have been shown to have higher sensitivities
and specificities than parasitological techniques. However, amplification targeting
repetitive sequences is more sensitive than the amplification of single copy sequences
(Masiga et al., 1992; Chappuis et al., 2005). Masiga et al. (1992) applied the PCR
technique to detect various trypanosome species by targeting repetitive DNA for
amplification. Using oligonucleotide primers designed to anneal specifically to the
satellite DNA monomer of each species/subgroup, this technique is able to accurately
identify T. simiae, three subgroups of T. congolense, T. brucei and T. vivax, and using a set
of more than one primer, this technique can detect mixed infections without use of
radioisotopes. This method uses crude preparations of template DNA, making this
technique rapid and ideal for large scale epidemiological studies. However, this technique
requires qualified personnel and a constant supply of electricity, resources which are
limited in most endemic regions (Masiga et al., 1992). Amplification of DNA by PCR is
able to accurately identify different trypanosome species and subspecies, enabling the
detection of mixed infections.
32
Recently, the Loop Mediated Amplification (LAMP) technique has been used in the
diagnosis of HAT (Njiru et al., 2008; Matovu et al., 2010). LAMP technique, developed
in 2000 by Notomi and colleagues is a DNA amplification technique which utilizes Bst
DNA polymerase with strand displacement activity and a set of four – six primers specific
to the template DNA being targeted. An inner primer containing sequences of the sense
and antisense strands initiates the amplification, and in an hour the target DNA is amplified
to 109 copies with high sensitivity. A LAMP-based test by Kuboki et al. (2003) to detect
T. b. brucei, T. b. gambiense, T. b. rhodesiense, T. evansi and T. congolense extracted DNA
showed that LAMP was 100 times more sensitive than PCR, and in vivo studies on T. b.
gambiense infected mice showed applicability of this test for disease diagnosis both in
reference laboratories and in field settings. Thekisoe et al.(2007) designed primers
targeting the 5.8S-rRNA-ITS2 sequence specific for T. b. gambiense, 18S rRNA targeting
T. congolense and T. cruzi, and a VSG specific for T. evansi (VSG Rotat 1.2). These
primers can detect 0.01 trypanosomes (1 femtogram trypanosome DNA), and the
technique is rapid and simple, showing applicability in field settings (Thekisoe et al.,
2007). In 2010, LAMP targeting random inserted mobile element (RIME-LAMP) for T.
b. gambiense detection was evaluated, and showed higher sensitivity and specificity
compared to TgSGP-PCR (Matovu et al., 2010). The need for four to six primers that
recognize six to eight regions of the target DNA makes it an expensive diagnostic test. In
addition, the need for trained personnel and power supply, make the approach inapplicable
in rural settings with limited or no health facilities.
Real time nucleic acid sequence-based amplification (NASBA/self sustained sequence
replication) targeting 18S rRNA of Trypanosoma brucei was developed for HAT diagnosis
in 2008 by Mugasa et al. (2008) NASBA is a RNA (or sometimes DNA) detection
technique that involves isothermal amplification of transcripts. It has been applied in the
detection of 18S rRNA of T. brucei (Mugasa et al., 2008). The test is sensitive, with a
detection limit of 10 parasites per mL. The ease and speed of detection of PCR and
NASBA products have been recently improved by the coupling of oligochromatography,
33
a technique whose sensitivity and specificity for T. b. gambiense and T. b. rhodesiense
differed, with the later recording low sensitivity and specificity on blood samples from
Uganda patients (Matovu et al., 2010).
The current techniques used in the diagnosis of HAT need improvement, and development
of a robust, cheap, simple, ready-to-use and large scale screening diagnostic kit is urgently
needed for T. b. rhodesiense detection as there is still no equivalent to the CATT for
population screening. An alternative approach to HAT diagnosis should be centred on the
detection of the parasite during its early stages of infection before the disease reaches its
late stages where treatment is both difficult and risky. This study evaluated the use of two
VSGs predominantly expressed during early stage of disease for early diagnosis of
Rhodesian HAT.
2.5 Disease Stage Determination
After pathogen detection in blood, it is necessary to determine the state of disease
progression by demonstration of parasites in the CSF or white blood cell counts in the
CSF, as recommended by the WHO (WHO, 1998). Stage determination and post treatment
follow up to determine cure or treatment failure are carried out by examination of the CSF
after patients undergo lumber puncture, a very painful and intrusive method that prevents
most people in endemic areas from being tested. According to WHO recommendations,
second-stage HAT is defined by the presence in the CSF of raised white blood cells (>5
cells/µl) and/or trypanosomes and increased protein content (>370 mg/liter, as measured
by the dye-binding protein assay) (WHO, 1998). The CSF white blood cell count is the
most widely used technique for stage determination (Chappuis et al., 2005), but the
threshold for treatment is controversial, differing from country to country (Table 2-1).
New born children (Sarff et al., 1976; Chappuis et al., 2005) and patients with bacterial,
viral or fungal meningitis however have high levels of wbc in the CSF, and this further
complicates the use of wbc counts in staging sleeping sickness disease.
34
Table 2-1. Different WBC/µL cut off for HAT staging in different sub Saharan
countries. Different countries apply different WBC/µL thresholds for HAT stage
determination, complicating the most commonly used method for stage determination and
subsequent treatment.
Etiological agent Country WBC/µL Reference
T. b. gambiense
Angola ≥20 Stanghellini and
Josenando, 2001
Cote d’Ivoire ≥20 Lejon et al., 2003
Uganda ≥20 Lejon et al., 2003
Sudan >5 Blum et al., 2006
Equatorial Guinea ≥10 Chappuis et al. 2005;
Blum et al., 2006
Central African
Republic
>5 Blum et al., 2006
Democratic Republic of
Congo
>5 Balasegaram et al. 2006;
Blum et al., 2006
Republic of Congo >5 Blum et al., 2006
T. b. rhodesiense
Malawi >5 WHO, 1998; Chappuis et
al., 2005
Uganda >5 WHO, 1998; Chappuis et
al., 2005
Detection of trypanosomes is carried out immediately after the lumber puncture because
the parasites lyse within 10 minutes in CSF. Demonstration of parasites in the CSF is
simple and cheap, but suffers from insufficient sensitivity, and is a very painful and
intrusive method that prevents most people in endemic areas from being tested or going
back for post treatment follow up.
35
Staging is also determined by assaying protein concentration in the CSF. In healthy
individuals, the CSF consists mainly of albumin (70%) and IgG (30%). In HAT patients
the protein concentrations are elevated from 15-60 mg/L to 100-2,000 mg/L. However,
protein concentrations can also be raised in first-stage illness due to diffusion in the CSF
of IgG which is present in high concentrations in the serum. This technique, though
simple, is limited by the difficulty in determining accurately the total protein concentration
in the CSF and is no longer used in the field and requires specific reagents which are more
often lacking in rural health centres (Chappuis et al., 2005). Demonstration of parasites in
the CSF and counting of white blood cells as methods of stage determination are limited
by low accuracy. In recent studies, biomarkers such as neopterin, CXCL-10, CXCL-13,
ICAM-1, VCAM-1 have been studied and shown potential for disease stage
determination, offering increased accuracy (Tiberti et al., 2013).
Figure 2-4. Lumber puncture to collect CSF for subsequent staging of sleeping
sickness. Patients with parasite in body fluids undergo lumber puncture, a process which
involves withdrawing cerebrospinal fluid (CSF) for analysis of trypanosome presence to
determine the stage of the disease. Image derived from FIND, 2012.
The CSF is subsequently analysed for presence of trypanosomes, while blood cell count
36
and protein quantities, allowing staging of the disease. Lumber puncture is a risky, painful
and intrusive method, with most patients showing that prevents most people in endemic
areas from being tested and/or going for follow-up to monitor treatment.
2.6 Antigenic Variation in African Trypanosomes
African trypanosomes are extracellular parasites that reside and replicate in the tissue
fluids and bloodstream of their mammalian hosts, where they are constantly exposed to
hosts immune system. Despite the continuous exposure to immune attack, trypanosomes
evade the immune response raised by the hosts’ immune system, leading to prolonged
pathological manifestations. This persistence in the mammalian host is due to a
phenomenon termed as antigenic variation, which involves switching the expression of
one variant surface glycoprotein (VSG) on the trypanosomes surface to another
immunogenically different VSG.
37
Figure 2-5. African trypanosome antigenic variation. Trypanosomes change the
expression of the surface coat, the variable surface glycoprotein (VSG) to an antigenically
distinct surface coat, enabling immune evasion. Image modified after Horn (2014).
The VSG is a dense cell surface coat that shields invariant surface antigens from immune
recognition. Antibodies against the expressed VSG kill the trypanosomes, but part of the
population survives due to stochastic switching at a rate of up to 10-2 to 10-7
switches/doubling time of 5-10 hrs (Turner & Barry, 1989). Three recombination
pathways which operate in hierarchy contribute to VSG switching, a hierarchy in which
telomeric VSGs are activated earliest in infections, followed by intact sub-telomeric array
VSGs, then lastly pseudogenes, which are recombined to form functional VSG mosaics
(Morrison et al., 2009). Gene conversion is a recombination pathway copying and
transferring a silent, functional VSG to the expression site (ES), replacing the resident
VSG. A second recombination pathway, the reciprocal telomeric VSG exchange involves
chromosome ends being exchanged by cross-over, moving a silent telomeric VSG into the
active ES and retaining the previously active VSG on the other chromosome, most likely
by double strand breaks (DSB) repair. The third recombination pathway involves the
generation of novel, functional VSGs by combining portions of the open reading frames
(ORFs) from at least two pseudogenes to generate VSG mosaics (Morrison et al., 2009).
38
VSG genes of T. brucei are transcribed in telomeric loci termed VSG expression sites
(ESs). The ESs active in bloodstream forms are polycistronic and contain several genes in
addition to the VSG, named ES-associated genes (ESAGs). So far 12 ESAGs have been
identified, some of which are present only in some ESs (Pays et al., 2001). The expressed
VSG is always located in an ES - found at the telomeres of the large and intermediate
chromosomes, while inactive VSG genes are scattered among different chromosomes
(Donelson, 2003). Each is a polycistronic unit, containing a number of ESAGs all
expressed along with the active VSG (Pays et al., 2001). While there are at least 20 known
expression sites, only a single one is ever active at one time. The RAD51 enzyme which
participates in DNA break repair and genetic exchange in many organisms is thought to
play a role in antigenic variation, but studies have shown that in its absence other
trypanosome factors provide for the VSGs switching (McCulloch & Barry, 1999).
2.7. The Variable Surface Glycoprotein (VSG)
The surface coat of bloodstream form trypanosome cell is composed of a 12 nm thick
glycoprotein known as the variable surface glycoprotein (VSG) (Vickerman, 1969).
Approximately 10 million molecules of the monolayer enwrap the trypanosome cell,
forming about 10-20% of the total bloodstream protein of the trypanosome cell (Bangs et
al., 1997). The VSG physically protects the underlying plasma membrane from
components of the hosts nonspecific immune effectors such as cytotoxic cells (Robinson
et al., 1999). The VSG molecule is approximately 60 kDa (Horn, 2014), and is anchored
to the plasma membrane by glycophosphatidylinositol (GPI). X-ray crystallography on
two thirds of the N terminal domains shows that the N terminals have a conserved rod like
shaped structures despite the different amino acid sequences, partly due to a conserved
disulphide bonds arrangement, indicating that VSGs share a similar back bone structure
from which emerge distinct epitopes from the different groups of amino acid side groups
(Donelson, 2003). The C terminal regions of the VSGs are conserved (Majumder et al.,
1981).
39
2.8 Sequential Expression of VSG
The majority of VSG genes are kept silent, and those expressed are sequentially switched
on (Ploeg et al., 1982). Even though there are >1000 VSG genes to select from, the genes
are expressed sequentially and in a hierarchy, a hierarchy in which telomeric VSGs are
activated earliest in infections, followed by intact sub-telomeric array VSGs, then lastly
pseudogenes, which are recombined to form functional VSG mosaics (Morrison et al.,
2009). At the beginning of the infective phase, VSGs are produced by parasites in the
insect salivary glands. A subset of the repertoire (about 12) of the VSGs which are
produced here remains during the first waves of parasitaemia in the mammal i.e. they
appear preferentially early in infection (Esser et al., 1982; Morrison et al., 2009) before
more complex genes predominate later on in the presence of a host-specific immune
response. The whole repertoire is then open to expression and there is preferred but not
fixed order of expression. VAT expression is reversible and hierarchical (i.e. non- random)
(Turner, 1999), and the possibility that there is an initial subset of VSGs that are expressed
sequentially during the early waves of parasitaemia gives hope for new methods of
diagnosis. Masiga and colleagues (unpublished work) carried out a study and identified
variable surface glycoproteins genes which where sequentially expressed predominantly
in all four vervet monkey models tested (see also Thuita et al., 2008). Two of the
determined genes formed the basis of this study, and were expressed in bacterial and insect
expression vectors. The recombinant proteins were serologically assayed against infected
monkey sera, and showed diagnostic potential by detecting anti CSG specific antibodies
in the samples assayed.
2.9. New serodiagnostic algorithms
Rapid diagnostic tests (RDTs) are fast, cheap, and easy to use on whole blood or plasma
and do not require electricity or a cold chain, making them applicable in screening
populations at risk under field conditions with limited resources. Furthermore, RDTs
provide an alternative to the card agglutination test for trypanosomiasis in T. b. gambiense
endemic foci. To improve specificity, synthetic and modified antigenic peptides (Van
40
Nieuwenhove et al., 2012) that will eliminate cross reactivity due to use of whole cell
lysates as in Gambian HAT kits can be used. Though this will mean increased kit costs,
generation of and simultaneous use of multiple antigens in rapid diagnostic tests (RDTs)
platforms are worth considering. For example, Büscher et al., (2013) developed HAT
Sero-Strip and HAT Sero-K-SeT tests specific for T. b. gambiense LiTat 1.3 and LiTat 1.5;
these were a dipstick and a lateral flow devices that cost about $2.50 each. Others include
SD BIOLINE HAT test by FIND and a prototype that uses recombinant ISG65 and native
VSG miTat 1.4 (sVSG117) developed by BBI Solutions (BBI) (Sullivan et al., 2014).
These attempts indicate potential for development of new and/or improved field
deployable sero-kits for Gambian HAT. The knowledge will be of immense importance in
diagnosis of the acute Rhodesian HAT.
41
CHAPTER THREE
3.0 MATERIALS AND METHODS
3.1 Generation of Recombinant VATs in Bacterial and Insect Expression Systems
3.1.1 Recovery of Cloned Variable Antigen Types (VATs)
Previously, Masiga and his colleagues (unpublished data) had recovered 21 VAT mRNA
from T. b. rhodesiense parasites during early onset of infections in vervet monkey models
(Cercopithecus aethiops) (see also Thuita et al., 2008). These VATs mRNAs were reverse
transcribed to cDNA, sequenced, cloned into pGEM-T Easy cloning vector (Promega,
Madison, USA) and subsequently sub-cloned into pRSET A expression vector
(Invitrogen, Carlsbad, USA). The recombinant plasmids were ultimately cryopreserved
by transforming competent DH5α and BL21 pLysS Escherichia coli (E. coli) cells
respectively. Two cryopreserved VAT gene isolates designated VAT3 and VAT4 were the
focus of this study. Of the identified VATs, these two VATs were shown to be the most
predominantly expressed in all vervet monkeys.
The recovery of the two cryopreserved VAT clones was carried out using Luria Bertani
(LB) (both agar and broth) selective media (Appendix 1). The cryopreserved stabilates
were aseptically streaked on freshly prepared LB agar plates with 100 µg/mL ampicillin,
and cultured overnight at 37 ºC in a bench top stationary incubator (Binder GmbhH,
Tuttlingen, Germany).
To confirm the presence of insert, colony PCR was carried out on randomly selected
colonies using Green Taq DNA polymerase (Genscript, Piscataway, USA). The reaction
mix consisted of 1 µL of 10 pmoles T7 universal primer and VAT3 and VAT4 reverse
primers (Appendix 2), 1 µl of 10× GenScript PCR Buffer, 0.4 µL of 1.25 mM MgCl2, 0.1
µL of 1 unit Taq DNA polymerase (GenScript Corporation, Piscataway, USA) then topped
to 10 µl with PCR grade water. Template DNA for PCR was a single bacterial colony for
42
each gene. Amplification was done using a programmed thermal cycler (PTC-100
Programmable Thermal Controller, MJ Research, Gaithersburg) at conditions which
included an initial denaturation/ activation step at 94 °C for 3 min, 35 cycles consisting of
denaturation at 94 °C for 30 sec, annealing at 54 °C for 1 min, and extension of 1 min 30
sec at 72 °C and a final extension step at 72 °C for 10 min. Following amplification, the
PCR products (5 µl) were gel electrophoresed on a 1% agarose gel for 1 hr at 7 V/cm and
visualized using the Kodak gel logic 200 imaging system.
3.1.2 Propagation and Purification of Recombinant Plasmids.
From the colony PCR, positive colonies for VAT3 and VAT4 each were cultured overnight
in 5 mL LB broth supplemented with 100 µg/mL ampicillin at 37 °C. The bacterial cell
cultures were cultured with vigorous shaking (150 rotations per minute) using the Environ
– Shaker, 3597-1 (Lab-line Instruments Inc, Conroe, USA). Subsequently, recombinant
plasmids were purified as described in the Quick-clean II Miniprep Plasmid Purification
Kit from GenScript (Piscataway, USA) with minor modifications. Briefly, the cells were
pelleted, resuspended in 250 µL resuspension buffer supplemented with RNAse A
solution, lysed and neutralized using 250 µL and 350 µL Lysis buffer and Neitralizing
buffers respectively. The plasmids were bound on the silica membrane, washed two times
with Wash buffer and eluted in 35 µL Elution buffer. Three (3) µL of the eluted plasmids
were electrophoresed as before (section 3.2).
Sequencing of the recombinant plasmids was outsourced from Macrogen Inc, Seoul,
South Korea and analysed using BioEdit Sequence Alignment Editor (Hall, 1999). From
the consensus sequences, bioinformatics analyses of the gene sequences were performed
using BLASTn, BLASTp and ExPASy Translate Tools (Gasteiger et al., 2003).
3.1.3 Cloning and Expression of VATs in Bacterial Cells
3.1.3.1. Amplification of VATs
The bacterial expression systems of choice were pRSET (Invitrogen, Carlsbad, USA) and
43
pBAD/His expression system. pBAD expression system was chosen after limited
expression success using the pRSET-his expression system. VAT4 full gene and VAT4
gene lacking the signal sequence (designated F0 and F1, respectively) were cloned into
pRSET B. VSG3 and VSG4 full genes were cloned into pBAD/His A expression.
The genes were amplified in a 50 µL PCR reaction volume consisting of 34.5 µl PCR
grade water, 10 µl of 5X Phusion HF Buffer, 1 µL of 10 mM dNTPs (final concentration
of 200 µM each), 1.5 µL of 10 pmoles forward and reverse primers (Appendix 2) (0.5µM
final concentration), 1 µL of template DNA (sequenced recombinant plasmids) and 0.5 µL
of 1 unit Phusion High Fidelity DNA polymerase (Thermo Scientific) to a final
concentration of 0.02 U/µl. Step-up PCR was used to amplify the VAT genes using the
PTC 100 Programmable Thermal Controller (MJ Research, Gaithersburg). The
amplification conditions included an initial denaturation/activation step at 98°C for 30
sec, followed by 40 cycles of denaturation at 98 °C for 45 sec, annealing at 46 °C for 1
min (first 20 cycles) and 50 °C (for the final 20 cycles), and an extension step at 72 °C for
1 min 30 sec, followed by a final extension step at 72 °C for 10 min. The PCR products
were electrophoresed as previously described and subsequently gel purified.
3.1.3.2 Gel Extraction of Amplified VAT Genes
Amplified VAT genes were excised and extracted as described in the QuickClean II Gel
Extraction Kit (GenScript, New Jersey, USA). Briefly, the excised gel slices were weighed
(not exceeding 40 mg) and 3 volume of binding buffer II to 1 volume of gel slice added.
The gel slices were dissolved by incubation at 55 °C for 6 min, and mixed with one volume
of research grade isopropanol to 1 volume of the gel slices. The extraction mixtures (750
µL) for each gene were transferred to separate spin columns, centrifuged at 6000 ×g for 1
min and flow through discarded. The DNA was bound to the matrix by adding of Binding
buffer II (500 µL) to the spin columns, centrifuged at 12000 ×g for 1 min, and the flow
through discarded. All subsequent centrifugations were carried out at this speed. The
columns were washed by 500 µL wash buffer and centrifuged as before and the flow
44
though discarded. The spin columns were centrifuged for an additional 1 min to remove
residual wash buffer, and the spin column transferred to sterile 1.5 microcentrifuge tubes.
The genes were eluted using 25 µL of elution buffer, and 3 µL electrophoresed as
previously described (section 3.2.1).
3.1.3.3 Cloning of amplified VAT genes into pGEM-T Easy Cloning Vector
pGEM-T Easy cloning system (Promega, Madison, USA) utilizes A/T- tail cloning
strategy. Since Phusion High Fidelity DNA polymerase does not add an adenine (A)
residue, polyadenylation of the gel purified PCR products was carried out using Green
Taq DNA polymerase enzyme (GenScript). A reaction mix consisting of 7 µL of each gel
purified product, 1 µL of 10× GenScript buffer, 1 µL of 2mM dATP and 1 µL (5 Units) of
Taq polymerase from GenScript was prepared, mixed gently and incubated at 70 °C for
30 min. A ligation reaction mix of 10 µL was prepared consisting of 3 µL of the A-tailed
PCR products, 5 µL of 2× Rapid ligation buffer, 1 µL pGEMT- Easy linear plasmids and
1 µL T4 DNA ligase. The ligation reactions were incubated overnight at 4 °C.
Five (5) µL of the ligation products for each gene were transferred into prechilled sterile
1.5 mL microfuge tubes and 50 µL of freshly thawed chemo-competent DH5α E. coli cells
added, mixed gently by flicking, and incubated on ice for 20 min. The cells were heat
shocked on a water bath at 42 °C for 45 sec, placed on ice for 2 min and Supper optimal
broth with Catabolite repression (SOC) media (950 µL) (Appendix 1) added to the cells.
The cells were cultured with vigorous shaking at 150 rpm in a shaker incubator at 37 °C
(Environ – Shaker, 3597-1, Lab-line Instruments Inc, Conroe, USA) for 1 hr, and
aseptically plated on LB/X-gal/IPTG plates supplemented with 100 µg/mL ampicillin (see
appendix 1). The plates were incubated at 37 °C overnight in a stationary bench top
incubator. The pGEM-T Easy vectors contain T7 and SP6 DNA polymerase promoters
flanking a multiple cloning region within α-peptide coding region of the enzyme
galactosidase. Insertional inactivation of the α- peptide allows identification of
recombinants by blue - white screening on indicator plates. Recombinant colonies were
45
inferred by blue - white screening. Positive (white) colonies were grown overnight in 5
mL selective LB broth (supplemented with 100 µg/ mL ampicillin) with vigorous shaking
(150 rpm) at 37 °C. The recombinant pGEMT-Easy vectors were purified using the Quick
Clean II Plasmid Purification Kit from GenScript as previously described (see section
3.1.2) and 3µL of the purified plasmids analyzed in a 1% agarose. The recombinant
plasmids were sequenced before the inserts were sub cloned into the respective expression
vectors and analyzed for correct orientation using ExPASy Translate tool.
3.1.3.4 Sub-cloning into bacterial expression vectors
The purified recombinant cloning vectors (pGEMT Easy) and expression vectors (pBAD/
His A and pRSET B) were restricted using FastDigest restriction enzymes from Thermo
Fischer Scientific. Restriction digestions using XhoI and KpnI for the pBAD/ His A vector
and BamHI and XhoI for pRSET B expression vector were performed. The restriction
reactions consisted of 12 µL nuclease free water, 3 µL 10 × FastDigest Green buffer, 13
µL plasmid DNA (1-2 µg) and 2 µL of FastDigest restriction enzymes in a final volume
of 30 µL. The restriction components were thoroughly mixed by gentle flicking, spun
down and incubated for 1 hr at 37 °C in a PTC-100 Programmable Thermal Controller.
The enzymes were inactivated at 80 °C for 5 min and the digested products
electrophoresed on a 1.5% agarose gel at 7V/cm for 1.5 hr. The expression vectors
(pBAD/His A and pRSET B) and the VSGs inserts were gel purified as described in
section 3.1.3.2.
The ligation reactions were set up in a total reaction volume of 10 µL, consisting of 5 µL
of 2 × Rapid Ligation Buffer (Promega), 1 µL of double digested plasmids, 3 µL of double
restricted VSG genes and 1 µL of T4 DNA ligase. The ligation reactions were incubated
at 4 °C overnight. Recombinant plasmids were transformed into chemo-competent DH5α
E. coli cells, propagated, the plasmids purified, sequenced and analysed as previously
described. From the consensus sequences, the recombinant plasmid sequences were
analyzed using BLASTn, BLASTp and ExPASy Translate Tool.
46
3.1.3.5 Expression and Analysis of VATs in Bacterial Cells
Bacterial strains used for expression were TOP10 and BL21pLysS for expressing genes
cloned in pBAD and pRSET expression vectors respectively. TOP10 E. coli strain is able
to transport the inducer L- arabinose but not able to metabolize it, while BL21 strain is
able to transport the inducer isopropyl β-D-thiogalactosidase (IPTGT) inside the cells to
induce expression of genes cloned downstream of a strong T7 promoter. TOP10 chemo-
competent cells transformed with recombinant pBAD/His A and BL21pLysS chemo
competent cells transformed with pRSET-B were grown on LB agar supplemented with
100 µg/ mL ampicillin and 34 µg/ mL chloramphenical (for BL21pLysS) and grown
overnight at 37 °C on a bench top incubator. One positive colony for each was inoculated
in 2 ml Supper Optimal Broth (SOB) (Appendix 1) with 100 µg/ mL ampicillin. The starter
cultures were grown overnight with shaking at 150 rpm to an OD600 1-2.
pBAD mock (pilot) expression study was carried out by sub-culturing 0.1 mL of the
overnight culture into five 10 mL SOB fractions and grown at 37 °C with vigorous shaking
to an OD600= 0.5. One mL aliquot of cells was removed from each starter culture,
centrifuged at maximum speed in a micro centrifuge tube for 30 sec, and the supernatant
aspirated. The cell pellets were frozen at -20 °C, making the zero time point sample. The
5 fractions were induced with 10 fold serially diluted L-arabinose concentrations as shown
in table 3-1 below. The cells were grown at 37 °C for 5 hrs with vigorous shaking at 225
rpm in a shaker incubator, centrifuged at maximum speed and the supernatant aspirated.
Table 3-1. Serially diluted L-Arabinose for protein expression pilot studies
Tube Volume (mL) Stock solution Final concentration
1 0.1 0.002% 0.00002%
2 0.1 0.02% 0.0002%
3 0.1 0.2% 0.002%
4 0.1 2% 0.02%
5 0.1 20% 0.2%
47
Expression of recombinant VATs using pRSET B expression systems involved culturing
1 mL of the overnight culture, and the subsequent inoculation into 50 mL SOB. The
bacterial cells were grown at 37 °C with vigorous shaking (225 rpm) to an OD 600 nm of
0.4-0.6. Time 0 samples were collected and pelleted as previously described. The cells
were divided into two, with one fraction induced with 1 mM IPTG final concentration and
the other fraction used as the uninduced expression control. The cells were grown at 37 °C
with vigorous shaking at 225 rpm. One mL from the induced and uninduced samples were
collected after every hour and pelleted as before and stored at -20 °C.
Protein expression analysis was done by one-dimensional sodium dodecyl sulphate -
polyacrylamide gel electrophoresis (SDS-PAGE) which involves dissociation of proteins
into their individual polypeptide subunits and separation in accordance to the polypeptides
size through migration through the polyacrylamide gels. Briefly, the protein samples
previously collected were dissolved completely in 1× Laemmli sample buffer (see
appendix 3). The samples were incubated at 95 °C for 7 min, spun down and 10 µL
separated at 10 mA, 100 V on a 12% (wt/vol) polyacrylamide gel (Sambrook et al., 1987)
for 2 hrs using the FTV 100Y Fischer Scientific electrophoresis system. The gels were
fixed and stained with Coomassie Brilliant Blue staining solution (Appendix 3) and
destained for visualization using white a light source.
3.1.3.6 Determination of the Solubility of the Recombinant VATs
Expressed recombinant proteins can either be soluble in the bacterial cells’ cytoplasm or
they may accumulate as insoluble aggregates called inclusion bodies due to aberrant
folding. To determine the solubility of the recombinant VSG proteins expressed, the
expressed pelleted cells were lysed completely by resuspension in 100 µL of 20 mM
phosphate buffer (pH 7.2) (Appendix 3) and frozen in liquid nitrogen. The lysates were
thawed at 42°C in a water bath. The freeze-thaw cycle was repeated three additional times,
and the insoluble fractions pelleted in a microfuge for 10 min at maximum speed at 4°C.
The supernatants were transferred into fresh 1.5 mL microfuge tubes. To 100 µL of
48
supernatant sample, an equal volume of 2× Laemmli sample buffer was added, and the
pellet resuspended in 100 µL of 1× Laemmli sample buffer. The protein fractions were
prepared and analysed as previously described (section 3.1.3.5).
3.1.3.7 Protein Scale Up and Purification
Two hundred and fifty (250) mL of SOB was inoculated with 500 µL of recombinant E.
coli cells cultured overnight and grown to an OD600 nm 0.4-0.6 at 225 rpm. The cells were
induced with IPTG or L-Arabinose at concentrations and durations (in hours) determined
by the pilot studies, grown at 37 °C with vigorous shaking (225 rpm) and the cells pelleted
for 10 min at 3000 ×g. Purification was carried out as described in the ProBond TM
purification system (Invitrogen, Carlsbad, USA). Preparation of buffers used in the
purification of recombinant proteins is shown in Appendix 4. The pelleted cells were
resuspended in lysis buffer (Native binding buffer supplemented with 8mg Lysozyme
enzyme) and incubated for 30 min on ice. The cells were lysed by sonication at high
intensity using the Soniprep 150 MSE Ultrasonic disintegrator (MSE Ltd, London, UK)
for six 10 second bursts with 10 second cooling intervals between each burst. The lysates
were centrifuged at 3000 ×g for 15 min at 4°C and the supernatant transferred onto
prepared ProBond ™ resin. The proteins were bound for 1 hr at room temperature using
gentle agitation on a rotating wheel to keep the resin suspended in the lysate solution. The
columns were washed four times with native wash buffer and clamped in a vertical
position and the cap on the lower end snapped off. The proteins were eluted with 12 mL
native elution buffer and collected in 1 mL fractions. Twenty (20) µL of the collected
fractions were analyzed by one-dimensional SDS-PAGE as describe in section 3.1.3.5.
3.1.3.8 Dialysis and Quantification of Purified Recombinant VATs
Purified recombinant VSG proteins in 50 mM NaH2PO4, 0.5 M NaCl and 250 mM
imidazole elution buffer were dialyzed against 10 mM PBS, pH 7.2 and 50 % sterile
glycerol at 4 °C overnight in a ratio of 1 mL eluted VSG protein to 200 mL PBS-glycerol
dialysis buffer. The dialyzed recombinant proteins were stored at -80 °C.
49
Dialyzed recombinant VSG proteins were quantified according to the Pierce™ BCA
(Bicinconinic acid) protein assay kit (Pierce, Rockford, USA). Briefly, 2mg/mL bovine
albumin serum standard was diluted using a diluents buffer composed of 0.1 M PBS (pH
7.2) and 100% glycerol in a ratio of 1:1 v/v to a final concentration of 250, 125, 50, 25, 5
and 0 µg/mL. The BCA working reagent was prepared by mixing 50 parts of BCA reagent
A with 1 part of BCA. One hundred (100) µl of the standards and VSG protein samples
were transferred into sterile protein free test tubes, and 2 mL of the BCA working reagent
added to each tube, mixed and incubated for 30 min at 60 °C using a water bath for even
heat transfer. The tubes were cooled to room temperature and OD562nm measured using the
Shimadzu UV-Visible Spectrophotometer BioSpec-mini (Kyoto, Japan). The
concentrations of the recombinant VSG genes were determined using the standard curve.
3.1.4 Cloning and Expression of VATs in Insect Cells
3.1.4.1 Amplification of VATs
pIZ/V5-His (Invitrogen, Carlsbad, USA) was the insect expression vector of choice. It
utilizes Orgyia pseudotsugata Immediate Early promoter (OpIE2) from the multicapsid
nuclear polyhedrosis virus (OpMNPV). The early promoters utilize the hosts’ transcription
machinery and do not require viral factors for activation. The forward primer included a
Kozak translation initiation sequence (G/ANNATGG) while the reverse primer had an
enterokinase cleavage recognition sequence (GACGATGACGATAAG) (Appendix 2).
The amplification was done as described in section 3.2.1 with the exception of the
annealing temperature used (48 °C and 52 °C). The amplification products were
electrophoresed and gel purified as earlier described (see sections 3.1.3.1 and 3.1.3.2).
3.1.4.2 Cloning VAT genes into pGEM-T Easy cloning vector
The amplified genes were excised, A-tailed and ligated into pGEMT Easy cloning vector
as described in section 3.1.3.3. Chemo-competent DH5α cells were transformed by 5 µLs
of the ligation products as earlier described. Positive colonies confirmed by blue – white
screening were propagated as described and the recombinant plasmids purified and
50
sequenced. The sequences were analyzed using BioEdit Sequence Alignment Editor.
From the consensus sequences, the VAT sequences were analyzed using BLASTn,
BLASTp and ExPASy Translate Tool.
3.1.4.3 Sub-cloning of VAT genes into the pIZ/V5-his expression vector
The recombinant pGEMT-T Easy plasmids and pIZ/ V5-His expression vectors were
restricted as described in section 3.3.3 with FastDigest BamHI and FastDigest XhoI
restriction endonucleases. The restricted expression vector and VAT genes were gel
extracted and ligated as earlier described and the recombinant plasmids propagated,
purified and sequenced as earlier described.
3.1.4.4 Culturing Spodoptera frugiperda (Sf21) Insect Cells
Cryopreserved Sf21 cells (Invitrogen, Carlsbad, USA) in freezing medium (Appendix 5)
were cultured in a 25 cm2 culture flask with 4 mL complete TNM FH medium (Appendix
5) supplemented with penicillin-streptomycin at a final concentration of 100 U/mL and
100 µg/mL respectively. The cells were evenly distributed across the flasks surface and
viewed under an inverted microscope. The cells were allowed to attach for 45 min inside
a sterile 27 °C, non- humidified, non- CO2 incubator (Eppendorf- New Brunswick Galaxy
48 S incubators, Edison, USA). After attachment, the medium with floating non-adherent
cells was replaced with 5 mL of fresh complete TNM-HH medium supplemented with
penicillin-streptomycin. The cells were cultured at 27 °C until they reached mid log phase
at about 90% confluence (approximately 3.8 × 106 cells). The cells were scaled up to
approximately 1.1 × 107 cells in a 75 cm2 culture flask and grown to confluence
(approximately 1.1 × 107 cells).
3.1.4.5 Transient Expression in Insect Cells
The cells were sloughed off with fresh medium as described before, and 1.2 × 106 cells
transferred into each well of a 6 well 60 mm tissue culture dishes. The cells were incubated
in a 27 °C incubator for 45 min to allow the cells to fully attach to the bottom of the dishes
to form a monolayer of cells. The cells were subsequently viewed under an inverted
51
microscope to verify adherence. Transfection mixtures were prepared in sterile 1.5 mL
microcentrifuge tubes, and consisted of 1 mL Grace insect media, Unsupplemented, 2
µg/µL plasmid constructs (pIZ/V5-His, VSG 3-pIZ/V5-His, VSG 4-pIZ/V5-His and GFP
-pIZ/V5-His) and 20 µL Cellfectin II reagent (Invitrogen, Carlsbad, USA). The
transfection mixtures were gently mixed and incubated at room temperature for 15 min.
The medium from the cells was carefully removed without disrupting the monolayer, the
cells washed completely with Grace insect media, Unsupplemented, and each transfection
mixture added to a separate well. Three replicates for each transfection were included.
The culture dishes were incubated at room temperature for 5 hr on a side-to-side, rocking
platform at 2 side to side motions per min. Complete TNM-FH medium (2 mL) was added
to each well and the dishes placed in a humidified 27 °C incubator. The cells and medium
were collected separately at 48, 72 and 96 hrs post-transfection and assayed for expression
of the target recombinant VSG proteins and fluorescence.
3.1.4.6 Testing for Expression of recombinant VATs
Twenty (20) µL of cells transfected with pIZ/V5-His empty vector and pIZ/V5-His GFP
constructs were fixed onto glass slides using an equal volume of 4% formaldehyde. The
cells were viewed under Light-emitting Diode (LED) and fluorescence using the
automated iMIC microscope (Till Photonics, Gräfelfing, Germany) Phantom camera.
Cells and medium from wells containing Sf 21 cell lines transfected with pIZ/V5-His
empty vector, pIZ/V5-His – VAT3 and pIZ/V5-His - VAT4 constructs were harvested and
assayed for expression using one dimension 12% SDS-PAGE. The cells were resuspended
in 100 µL of 1X Laemmli sample buffer, while 100 µL of the medium was mixed with
100 µL of 2x Laemmli sample buffer. The samples were boiled for 10 min and 10 µL
loaded onto a 12% SDS polyacrylamide gel. Electrophoresis was carried out at 10 mA and
100 V as previously described. The gels were fixed and stained with Coomassie Brilliant
blue and destained to visualize the proteins.
52
3.1.4.7 Scale-up, Purification and Quantification of Recombinant VATs
Sf21 cells were cultured in three 75 cm2 culture flasks using Sf-900™ SFM to confluence.
The cells were transfected as before with 2µg pIZ/V5-His empty vector, pIZ/V5-His –
VAT3 and pIZ/V5-His - VAT4 constructs, and incubated at 27 °C for 96 hr. The cells and
the media were harvested separately. The cells were resuspended in 8 mL Native binding
buffer (Appendix 4), the cells lysed by two freeze thaw cycles and passed four times
through an 18-gauge needle and centrifuged at 3000 ×g for 15 min. Proteins in the medium
were dialyzed against Native binding buffer as well. The lysate and dialyzed secreted
proteins were bound for 30 min at room temperature on a rotating wheel. The resin was
settled by low speed centrifugation (800 ×g) and the supernatant discarded. The column
was washed and the proteins eluted as described before (section 3.3.5.4). Subsequently,
the eluted proteins were analyzed by one dimensional SDS PAGE, and subsequently
dialyzed against 0.1M PBS – 100% glycerol overnight as previously described and stored
at -80°C. Dialyzed recombinant VSG 3 and VSG 4 antigens were quantified using the
BCA protein assay as described in section 3.1.3.8.
3.2 Evaluation of the Diagnostic Potential of the Generated Recombinant VATs
3.2.1 Infection of Monkey disease models and sera collection
Previously, Thuita et al. (2008) had infected nine monkeys with four different T. b.
rhodesiense strains (with varying pathogenicity and collected from different disease foci
in Kenya and Uganda). Briefly, nine vervet monkey models (Chlorocebus aethiops) were
infected via a single bite of tsetse infected with T. b. rhodesiense. The monkeys were
randomly allocated to four experimental groups, and each group infected with a different
strain of T. b. rhodesiense as follows;
53
Table 3.2: A table showing the different T. b. rhodesiense strains used for infecting
monkeys belonging to four experimental groups, and their years of isolation.
T. b.
rhodesiense
strain
Geographic region
of strain isolation
Year of
strain
isolation
Monkeys per
experimental
groups
Monkey
tags
3741 Busoga, Kenya 2003 3 476,515,536
3804 Bukhayo, Kenya 1989 2 523, 579
3801 Busia, Kenya 1989 2 556,574
3804 Tororo, Uganda 1972 2 554,555
Blood was harvested from monkeys from all the experimental groups after the 3rd, 7th,
11th, 15th days post infection (dpi) representing early stage of disease and 19th dpi
representing onset of late stage disease. The collected blood was stored at -80°C. Twelve
sera samples available at icipe’s cryo-bank were used in this study, with each T. b.
rhodesiense strain represented. Sera collected from two uninfected monkeys were used as
controls in the immunoassays.
3.2.2 Serum and Immunodiffusion Gel Set-up
Archived monkey blood samples infected with T. b. rhodesiense were thawed on ice and
centrifuged at 1000 ×g for 15 min using the 5417R Eppendorf bench top centrifuge. The
serum was carefully aspirated from the pelleted erythrocyte cells and transferred to sterile
1.5 mL micro tubes. Sera from two uninfected monkeys were included as controls in this
study, and were prepared as described above.
One gram of agarose, molecular grade, was dissolved in 50 mL 0.1 M PBS (pH 7.2)
supplemented with 0.03% sodium azide (w/v), and dissolved completely by boiling, then
cooled to 55 °C. Three (3) g of polyethylene glycol (PEG) was dissolved in 50 mL 0.1 M
PBS (pH 7.2) with supplemented with 0.03% (w/v) sodium azide and dissolved
completely by warming it to 55 °C. The agarose and PEG solutions were mixed in equal
54
volumes to make a final concentration of 1% and 3% respectively, and the gel cast on
clean dry glass plates. Holes were punctured on the gel using a gel punch and a water
vacuum pump used to carefully aspirate the gel from the punctured holes.
3.2.3 Evaluation of Diagnostic Potential via Ouchterlony Immuno-assays
The expressed purified recombinant antigens (15 µL) were loaded in the central well and
the same volume of neat monkey serum loaded in the peripheral/ outer wells, separated
by a well of 15 µL 0.1 M PBS, pH 7.2. Negative controls of monkey sera collected from
none-infected monkeys were also assayed. Care was taken to avoid over filling or under
filling the wells. The agarose plates were placed in moist air tight plastic containers, and
incubated for 24-48 hrs at 37°C. The formation of white precipitin lines was monitored by
viewing the plates in a dark field light viewing box. For permanent results, the plates were
stained using Coomassie brilliant blue after rinsing the plates overnight with 0.1 M PBS,
pH 7.2 and drying/ pressing the agarose overnight. The plates were viewed and
photographed.
55
CHAPTER FOUR
4.0 RESULTS
4.1 Generation of Recombinant VATs in Bacterial and Insect Expression Systems
4.1.1 Expression of Recombinant VATs in Bacterial Cells
The amplification of VSG genes from recovered cryopreserved bacterial colonies by
colony PCR showed DNA bands of approximately 1.5 kb product for both VAT3 and VAT4
in a 1% ethidium bromide stained agarose gel (Figure 4-1A), confirming presence of these
genes in the cryopreserved bacterial stabilates. Sequence analysis confirmed the sizes of
the VSG genes to be 1452 and 1449 basepairs respectively. BLASTn and BLASTp
resulted in hits closely similar to trypanosome VSGs. Amplification of VAT3 and VATs4
genes with primers having restriction sites to enable cloning of the genes into bacterial
expression vectors (both pRSET and pBAD) gave approximately 1.5 kb for full genes
(Figure 4-1B), while amplification of VSG 4 without the signal peptide sequence yielded
a 1382 bp product. After cloning, restriction of recombinant pGEMT-Easy and pRSET B
plasmids showed two bands at approximately 3 kb and 1.5 kb for the linearized pGEMT
Easy/pRSET B plasmids and VSG genes respectively (Figure 4-1C). Sequence analysis
by ExPASy translate tool confirmed successful cloning of VAT3 and VAT4 genes into
pBAD/His A at XhoI and KpnI and pRSET/His B at BamHI and XhoI sites. Moreover, the
genes were shown to be cloned in frame with the N terminal six histidine tags (Figure 4-
1D), necessary for subsequent purification after expression of the recombinant proteins.
The gene sequence results of VATs will not be presented here, because Masiga and
colleagues, still have interests in these sequences.
56
Figure 4-1: Results of VAT3 and VAT4 amplification, cloning and bioinformatics
analysis. (A) Amplification of cryopreserved bacterial stabilates by colony PCR and (B)
Subsequent amplification with primers for cloning into pBAD or pRSET bacterial
expression vectors produced 1.5 kb products. (C). Restriction digestion of recombinant
pGEMT Easy and pRSET/ His B vectors after cloning. (D) A schematic representation of
recombinant bacterial expression vectors after sequence analysis, showing successful
cloning of the genes in frame with the N- terminal 6 his tag.
Recombinant VATs were successfully expressed after transformation of competent E. coli
cells and induction with 1M IPTG or 0.02% L-Arabinose for pRSET B and pBAD/His A
57
expression vectors respectively and grown for 4 hrs post induction, as shown in figure 4-
2 below. Purification of the recombinant 6× his-tagged VSG 3 and VSG 4 proteins
expressed using pRSET and pBAD expression vectors using immobilized metal affinity
chromatography was also successful as shown in panels (B) and (C) respectively.
Figure 4-2: SDS-PAGE analysis of expression and purification of recombinant VAT3
and VAT4 in bacterial cells. (A) Recombinant VAT4 (full gene) expressed in pRSET B
bacterial expression system gave a product of approximately 55 kDa. (B) VAT4 without
the signal peptide, (F1), expressed using pRSET B expression system gave a product of
about 45 kDa. (C) Full length VAT3 recombinant was expressed using pBAD/His A
expression system, similarly giving a product at about 55 kDa. U, Uninduced total cell
lysate control; I- Induced total cell lysate; SF, Soluble fraction; IF, Induced insoluble
fraction; P- Purified recombinant VSG protein.
4.1.2 Expression of Recombinant VATs in Insect Cells
Amplification of VAT3 and VAT4 genes with primers for cloning into BamHI and XhoI
sites of the pIZ/V5-His insect expression vectors were successful, as confirmed by agarose
gel electrophoresis of amplified products, restriction of the recombinant plasmids with
BamHI and XhoI restriction endonucleases, and subsequent sequence analysis (Figure 4-
3). Analysis of the sequenced recombinant plasmids by ExPasy Translate Tool confirmed
that the two genes were in their correct ORFs and had been cloned in frame with the C
terminal histidine tag.
58
Figure 4-3: Agarose gel results of VAT3 and VAT4 gene amplification and
bioinformatics results of recombinant pIZ/V5-his constructs. (A). Amplification of
VAT genes produced approximately 1.5 kb DNA products. (B). Restriction digestion of
recombinant pIZ/V5-his VAT3 and pIZ/V5-his VAT4 constructs produced two bands at 3
kb and 1.5 kb, representing vector DNA and VSG DNA inserts respectively. (C) A
schematic representation of recombinant pIZ/V5-His – VSG 3 or pIZ/V5-His – VSG 4
constructs after sequence analysis.
A green fluorescent protein gene (GFP) cloned into pIZ/V5-His expression vector
included as a positive control for expression was analyzed for both transfection efficiency
and expression in Sf 21 cell lines. The images of Sf 21 cells transfected with pIZ/V5-His
– GFP recombinant construct were compared to images of Sf 21 cells transfected with an
empty pIZ/ V5-His expression vector (Figure 4-4A). The cells transfected with an empty
59
expression vector did not emit fluorescence under UV light at 48, 72 and 96 hrs post
transfection. Cells transfected with a recombinant pIZ/V5-His carrying the GFP gene
fluoresced under UV light, with the fluorescence increasing with time.
Sf 21 cells transfected with recombinant pIZ/V5-His – VSG 3 and pIZ/V5-His – VSG 4
constructs and pIZ/V5-His only control by lipid mediated transfection, harvested after 48,
72 and 96 hrs post transfection and analyzed for expression by SDS-PAGE, and showed
that VSG proteins were expressed and extracellularly secreted into the growth media
(Figure 4-4B). The recombinant proteins were approximately 55 kDa. Media from cells
transfected with pIZ/V5-His only control did not show the prominent band at 55 kDa.
60
Figure 4-4. Microscopy and SDS PAGE results of recombinant protein expression in
insect cells (Sf21 cell line. (A). Sf 21 cells transfected with pIZ/V5-His – GFP
recombinant construct showed fluorescence under UV light, illustrating successful
transfection and expression of the green fluorescent protein using the pIZ/V5-his insect
expression vector. (B) Expression of VAT3 and VAT4 recombinant antigens showed a 55
kDa protein in the secreted fraction. Protein ladder is shown next to the gel image. IPs-
intracellular proteins, EPs-Extracellular proteins.
61
4.2 Evaluation of the Diagnostic Potential of the Expressed Recombinant VATs
4.2.1 Serodiagnostic Potential of Recombinant VAT3 and VAT4
Following expression and purification, recombinant VAT3 and VAT4 were dialyzed and
serologically assayed against archived monkey serum infected with T. b. rhodesiense.
Twelve monkey sera samples infected with four different T. b. rhodesiense strains
collected 3, 7, 11, 15 and 19 days post infection were assayed against the recombinant
VATs expressed in bacterial and insect expression systems. Formation of precipitin lines
of immuno-recognition between the expressed recombinant variable surface antigens and
anti VSG specific antibodies in the sera samples was observed as shown in figure 4-5.
Sera collected from two uninfected healthy monkeys did not form immuno-precipitin
lines, illustrating the specificity of the recombinant antigens.
From the immunoassays, the detection rates (in percentage) were calculated for the
recombinant VATs against the assayed infected monkey sera by dividing the number of
times the recombinant VATs formed precipitin lines of immunorecognition by the total
number of infected sera samples assayed then multiplied the ratio by 100. Recombinant
VAT3 and VAT4 expressed in bacterial cells had detection rates of 88% and 0%
respectively, while VAT3 and VAT4 expressed in insect cells had detection rates of 97%
and 83% respectively.
62
Figure 4-5. Ouchterlony double diffusion results showing immune recognition of
recombinant VATs and sera collected from T. b. rhodesiense infected monkeys.
Precipitin lines formed between (A) recombinant VAT3 expressed in bacterial cells and
(B) recombinant VAT4 expressed in insect cells against sera collected from infected
monkeys (C) Precipitin lines did not form when sera collected from uninfected monkey
was reacted against VAT3 expressed in bacterial cells (upper pattern), while the same
antigen formed precipitin lines against sera collected from an infected monkey (lower
pattern).
4.2.2 Potential Application of VATs for Early Disease Detection
Twelve monkey sera samples infected with four different T. b. rhodesiense strains
collected 3, 7, 11, 15 and 19 days post infection were assayed against the recombinant
VATs expressed in bacterial and insect expression systems. The serological assays were
repeated three times. The formation of a white precipitin lines, indicating immuno-
recognition between the expressed antigens and anti-VSG specific antibodies, were
observed after a 24 hr incubation, and are represented by (+) in Table 4-1 below. Absence
of immuno-recognition is represented by (-). Sera collected from uninfected monkeys did
not form precipitin lines with all the expressed recombinant antigens.
The sera samples assayed were collected 3, 7, 11 and 15 dpi (representing early stage
disease) and 19 dpi (representing onset of late stage disease) from monkey models infected
by the bite of a single infected tsetse fly to mimic the natural infection route in man. Four
63
strains of T. b. rhodesiense collected from different geographic regions in Uganda and
Kenya were used to infect the monkey models (Thuita et al., 2008). Precipitin lines were
formed between VSG specific antibodies and the expressed recombinant VATs from sera
samples collected as early as 3 dpi through to the 15th dpi representing the early
stage/haemolymphatic stage of the disease, and 19 dpi when the parasite had crossed into
the CNS (Table 4-1).
64
Table 4-1: Diagnostic performance of recombinant VATs against monkey sera samples infected with four different T.
brucei rhodesiense KETRI strains collected at different days post infection (dpi). Uninfected sera controls are also shown.
Each immunoassay was repeated three times. + represents immunorecognition between recombinant VATs and anti-
trypanosome specific antibodies in the serum, while – represents absence of immunorecognition for each experiment.
Monkey
experimental
group
T. b. rhodesiense
strain
Geographic
region of strain
isolation
Days post
infection
(dpi)
Bacterial expression system
VSG 3 VSG 4
Insect expression system
VSG 3 VSG 4
552 KETRI 3741 Busoga, Kenya 3 +++ --- +++ ++-
554 KETRI 3928 Tororo, Uganda 3 +++ --- +++ +++
554 KETRI 3928 7 +-+ --- +++ +++
555 KETRI 3928 3 +++ --- +++ -++
555 KETRI 3928 7 +-+ --- +++ +-+
555 KETRI 3928 11 +++ --- +++ +++
555 KETRI 3928 15 +++ --- +++ +++
555 KETRI 3928 19 -++ --- -++ -++
556 KETRI 3804 Bukhayo West,
Kenya
3 +++ --- +++ +++
574 KETRI 3804 3 -++ --- +++ +++
574 KETRI 3804 7 +++ --- +++ +-+
657 KETRI 3801 Busia, Kenya. 7 +++ --- +++ +++
716 Uninfected N/A --- --- --- ---
720 Uninfected N/A --- --- --- ---
70
CHAPTER FIVE
5.0 DISCUSSION
5.1 Generation of Diagnostic Recombinant VATs in Bacterial and Insect Expression
systems
Recombinant VAT3 and VAT4 were successfully generated in bacterial and insect cells
and subsequently purified as histidine tagged fusion proteins (the yields are shown in
Appendix 6, Table A-2). The VATs expressed from full length VSG genes were of the
same molecular weight irrespective of the expression systems, suggesting that post
translational modification possibly did not occur in insect cells. Expression in bacterial
system was fast (between 3–4 hours post induction), reproducible and used inexpensive
culture medium. On the contrary, expression in insect cells was time consuming (3-4
weeks of cell culture followed by 48-96 hrs post transfection) and used expensive culture
medium. Overall comparison of the two expression systems however indicates that
expression in insect expression system results in more recombinant protein yields and an
easier purification procedure. Removal of the fusion tag from the purified recombinant
proteins is possible using enterokinase enzyme, however immuno-recognition of the 6
histidine tagged VATs by antibodies in the sera samples assayed suggests that the histidine
tags (both N and C terminally placed) do not alter epitope conformations during folding
of the proteins in the two expression systems.
A VAT diagnostic tool based on recombinant antigens offers advantages over the use of
native antigens. For example, the CATT/T. b. gambiense mass screening tool is based on
natively produced LiTat 1.3 VSG antigen. The diagnostic antigen is mass produced by
infecting laboratory rodents using human infective T. b. gambiense parasites expressing
LiTat 1.3 VSG. The use of recombinant antigens eliminates the need to culture human
infective trypanosomes. Moreover, generation of recombinant antigens is cheaper and
easy to standardize, ultimately leading to affordable diagnostic tools (Rogé et al., 2014).
71
Several diagnostic antigens have been expressed in heterologous expression systems and
successfully applied in diagnosis of human diseases, further increasing confidence in the
recombinant antigens assayed in this study. For instance, a fungal disease termed
paracoccidioidomycosis (PMC) caused by Paracoccidioides brasiliensis, has been shown
to be diagnosed using an exocellular glycoprotein gp43 expressed in E. coli cells (Diniz
et al., 2002). Insect expression systems have also been used in the generation of diagnostic
antigens. In 2001, Urakawa and colleagues expressed a variable surface glycoprotein
specific to T. evansi in Sf 21 cells, and showed their potential in diagnosis of Surra disease
in livestock.
5.2 VATs Have Diagnostic Potential for Antibody Detection
This study set out to evaluate the diagnostic potential of two VSGs predominantly
expressed during early stages of T. b. rhodesiense infection as potential candidates for
diagnosis. Immunorecognition of the purified recombinant antigens by anti-VSG specific
antibodies in monkey sera infected with T. b. rhodesiense illustrated the diagnostic
potential of the recombinant antigens assayed (Figure 4-5; Table 4-1). The interaction is
specific between VAT epitopes and anti VSG specific antibodies, and results in the
formation of precipitin lines. Recombinant VAT4 expressed in bacterial cells did not react
with the sera samples assayed, probably due to low yields (see Table A-2), or aberrant
folding of the recombinant VAT. Non reactivity of expressed recombinant VATs with sera
from uninfected infected monkey controls indicates the specificity of these antigens.
Monkey models have been shown to mimic HAT clinically, immunologically and
pathologically (Schmidt & Sayer, 1982), validating the use of sera from vervet monkeys
in this study as a model for HAT disease. The ability of the expressed recombinant antigens
to be immunologically recognized by antibodies from T. b. rhodesiense infected monkeys
gives confidence that they can be potent diagnostic markers for screening specific anti
VAT3 and anti VAT4 antibodies in human serum samples. Furthermore, the two VSGs
assayed were shown to be predominant in the four different strains of T. b. rhodesiense
72
isolated from different locations, increasing the geographic user range of the VSG
diagnostic candidates.
The confirmation of diagnostic potential of the individual recombinant antigens offers a
valuable alternative to parasitological demonstration of parasites in body fluids, the
recommended diagnostic gold standard by WHO, which suffers from low sensitivity and
specificity, leading to approximately 30% of individuals being missed (Chappuis et al.,
2005). Even though antibody detection tools do not offer direct evidence for the presence
of trypanosomes, a VAT based diagnostic tool able to detect antibodies elicited upon
contact with the parasite could form more potent diagnostic tools for disease detection
compared to parasitological tools which are in addition not amenable to mass screening.
The ability of expressed recombinant VATs to show immuno-reactivity with sera collected
three days post infection (dpi) up to 15 dpi demonstrate potential application in early
diagnosis of sleeping sickness caused by T. b. rhodesiense. The mean peak parasitaemia
in the monkey models used was 3-4 days, with an overall median of parasite penetration
into the CSF at 16 dpi (Thuita et al., 2008). Early treatment of sleeping sickness caused
by T. b. rhodesiense is essential since treatment of late stage disease is risky and
complicated due to the toxicity of Melarsoprol, the only drug option available (Kennedy,
2013). The assayed recombinant VATs show promise as ideal diagnostic candidates for
early disease detection. Moreover, detection of specific VSG 3 and VSG 4 antibodies at
19 days, which represents the late stage of the disease when the parasite has crossed into
the CNS, suggests potential application of assayed VATs in diagnosis of sleeping sickness
during both stages of the disease. However, antibody detection tests are limited by the fact
that as antibodies remain in circulation days after their production by host’s immune
system, detection of circulating antibodies instead of antigens could lead to false positives
after treatment or self cure. One way of resolving this limitation would be combing the
VSGs with other diagnostic candidates such as ISG 65 and ISG 75 (Ziegelbauersq &
Overaths, 1992). These invariant surface glycoproteins have been reported to be highly
immunogenic, eliciting production of specific antibodies, and are potent diagnostic
73
molecules for sleeping sickness (Sullivan et al., 2013; Sternberg et al., 2014).
Parasitological tests that also demonstrate parasites during active infection can be applied
following screening to confirm and/or discriminate active infection from cured cases.
In the absence of a serological tool for screening populations at risk and animal reservoirs
of T. b. rhodesiense, this study describes the potential application of recombinant VSG
antigens applicable in distinguishing mixed infections, especially in Uganda in
combination with the CATT/T. b. gambiense mass population screening tool. Currently,
no equivalent of the CATT/ T. b. gambiense serological screening diagnostic tool is
available for screening populations at risk of T. b. rhodesiense infection (Chappuis et al.,
2005). The potential convergence of Gambian and Rhodesian HAT in Uganda (Picozzi et
al., 2005) complicates diagnosis, treatment and epidemiological studies of the disease, and
the introduction of a sensitive, robust and field applicable serodiagnostic tool for screening
T. b. rhodesiense infections would facilitate early and accurate diagnosis and thus safe and
correct treatment in foci where the two form of the disease occur. The CATT/T. b.
gambiense and LATEX/T. b. gambiense are serological tests based on lyophilized
trypanosomes expressing LiTat 1.3 and purified LiTat 1.3, 1.5 and 1.6 variable surface
antigens, respectively. These are predominantly expressed VSGs which agglutinate with
T. b. gambiense specific antibodies. A VAT – antibody test for T. b. rhodesiense based on
predominant variable surface antigens could form the basis for development of a similar
serodiagnostic tool to screen both humans and animal reservoirs of sleeping sickness, and
improve surveillance and epidemiological studies impacting control strategies.
Serological assays are cheap, amenable to field conditions and do not require material
resources.
74
CHAPTER SIX
6.0 CONCLUSIONS AND RECOMENDATIONS
6.1 CONCLUSIONS
This study delivers proof of concept that assayed VAT recombinant antigens
predominantly expressed during early onset of T. b. rhodesiense infection have diagnostic
potential. The diagnostic potential of each separate recombinant VAT was confirmed by
ouchterlony double immunodiffusion test on sera collected from monkeys infected with
different T. b. rhodesiense strains and two negative sera samples. VSG3 and VSG4
recombinant antigens could form the basis for development of a simple, cheap, robust and
sensitive diagnostic tool applicable in the field setting. Moreover, detection of the
expressed variable surface antigens by antisera collected at 3-15 dpi from monkey models
infected with different T. b. rhodesiense strains suggests that these antigens have potential
to be important candidates for consideration in development of a novel diagnostic tool for
early disease detection. However, further evaluation and testing is required with a large
panel of HAT positive and negative sera collected from endemic and non endemic areas,
and sera collected from patients suffering from other infections which co-exist in sub
Saharan Africa such as malaria and Leishmaniasis to assay for cross reactivity.
6.2 RECOMMENDATIONS
This study suggests that recombinant VAT expressed predominantly by T. b. rhodesiense
have diagnostic potential for early case disease detection as shown by ouchterlony test, a
simple serological test. Moving forward, a larger panel of VSGs expressed predominantly
during early stages of infection should be assayed, both as single antigens and/or in
combination, to form a more robust panel of antigens which could be included in
combination to develop a serodiagnostic tool. This will improve the geographic user range
for this test, as different strains of T. b. rhodesiense exist (Barry & McCulloch, 2001).
Secondly, screening the recombinant antigens with a large panel of human sera from
75
endemic and non-endemic areas needs to be carried out, with a more sensitive serological
test such as ELISA before development and deployment of rapid kits based on these
antigens. This will enable the determination of the sensitivity, specificity with the current
diagnostic tools currently available for the diagnosis of T. b. rhodesiense sleeping
sickness.
Thirdly, the diagnostic performance of the assayed variable surface antigens could be
improved by testing them in combination with other highly immunogenic trypanosome
antigens. Two predominantly expressed invariant surface glycoproteins, ISG65 and
ISG75, expressed specifically in bloodstream form trypanosomes (Ziegelbauersq and
Overaths, 1992), form good combination antigens to improve the sensitivities and
specificities of the two assayed variable surface antigens. ISG65 and ISG75 have been
used in combination for diagnosis of Gambian HAT (Sternberg et al., 2014), and could be
used in combination with the assayed recombinant antigens in this study to form a basis
for the development of a cheap, simple, robust and field applicable diagnostic tool for
diagnosis of T. b. rhodesiense HAT.
76
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APPENDICES
Appendix 1: Bacterial Culture and Transformation media
a) Isopropyl β-D-1-thiogalactopyranoside (IPTG) stock solution 0.1M contains
1.2g IPTG dissolved in 50ml distilled water and stored at 4°C
b) 5-bromo-4-chloro-3-indolyl-β-D-galactopyranoside (X-Gal) 2ml was
prepared by dissolving 100g of 5-bromo 4-chloro 3-indolyl β-D-galactoside in 2ml of N,
N'-dimethyl-formamide. It is stored at -20°C in an aluminium foil covered container.
c) Luria-Bertani (LB) medium: 10g Bacto-tryptone, 5g Bacto-yeast extract and
5g NaCl was dissolved in a litre of distilled water and its pH adjusted to pH 7 using NaOH.
This was then autoclaved before use.
d) LB plates with ampicillin, IPTG and X-Gal: 15g agar was added to a litre of
LB medium then autoclaved. This was allowed to cool to 50°C before ampicillin was
added to a final concentration of 100μg/ml.
e) 2M Mg2+ stock: Prepared by mixing 20.3g MgCl2 •6H2O and 24.65g MgSO4
•7H2O and distilled water to a final volume of 100ml then filter sterilized.
f) Supper Optimal Broth: 100 mL was prepared by mixing 2g of bacto-tryptome,
0.5g bacto-yeast extract, 1ml 1M NaCl, 0.25ml 1M KCl, 1ml 2M Mg2+ stock (filter
sterilized) and distilled water to a final volume of 100ml.
g) Super Optimal broth with Catabolite repression (SOC) medium: 100ml was
prepared by mixing 2g of bacto-tryptome, 0.5g bacto-yeast extract, 1ml 1M NaCl, 0.25ml
1M KCl, 1ml 2M Mg2+ stock (filter sterilized), 1ml 2M glucose (filter sterilized) and
distilled water to a final volume of 100ml.
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Appendix 2: Primer sequences
Table A-1: Sequences of primers used to amplify VAT3 and VAT4 genes for
subsequent cloning into pRSET, pBAD and pIZ/V5-His expression system in frame
with the 6-his tag
Gene and Vector Primer Sequence (5' to 3')
T7 Promoter TAATACGACTCACTATAGGG
VSG 3-pBAD/His A
For: GTGCTCGAGATGCGGCCCACCACTTTAG
Rev: GGCGGTACCATTAAAAAAGCAAAACTGC
VAT4-pBAD/His A For: GTGCTCGAGATGCGGCCCACCACTTTAG
Rev: GGCGGTACCATTAAAAAAGCAAAACTGC
VAT4-pRSET B (F0) For: CAGGATCCGATGCGGCCCACCAC
VAT4-pRSET B (F1) For:GCGGATCCAATCACACAACCGTGTGAGGAAG
Rev: ACTCGAGCCATTAAAAAAGCAAAACTGC
VAT3-pIZ/V5-His
For: GCGGGATCCATGCGGCCCACCACTTTAGC
Rev:GCCTCGAGCGATCCTTATCGTCATCGTCAAAAAGC
AAAACTGCAAGCC
VAT4-pIZ/V5-His
For: GCGGGATCCATGCGGCCCACCACTTTAGC
Rev:CCTCGAGCGATCCTTATCGTCATCGTCAAAAAGCA
AAACTGCAAGCC
Appendix 3: SDS-PAGE Buffers
Electrophoresis/Running Buffer: 1000mL was prepared by mixing 15.10 g Tris base, 94 g
of Glycine and 5.0g SDS. The pH was 8.3 without adjusting.
a) Laemmli sample buffer: 10 mL 4×stock solution was prepared by mixing 2
mL 0.2M Tris HCl, pH 6.8, 0.8g SDS, 4 mL 100% glycerol and 8 mg bromophenol blue
and 0.4 mL 14.7M β- mercaptoethanol to make 50 mM Tris HCL, 2% SDS, 10% glycerol
, 0.002% bromophenol blue and 1% β- mercaptoethanol.
b) Separating buffer: To make 100 mL 1.5M Tris HCl, 18.15g of Tris base was
99
dissolved in 90 mL H2O, pH adjusted to 8.8 then topped to 100 mL.
c) Stacking gel buffer: To make 100 mL of 5 × 1M Tris HCl, 12.114g of Tris
base was dissolved in 90 mL H20, the pH adjusted to 6.8 and topped to 100 mL.
d) Coomassie brilliant blue staining solution: To prepare 100 mL, 50 mL H2O
was mixed with 40 mL methanol and 10 mL Glacial Acetic Acid. 0.2g Coomassie brilliant
blue was dissolved completely and the staining solution filtered using a whatmann filter
paper.
e) Phosphate Buffered Saline (PBS): To prepare 100 mL of 1× PBS, 8g of NaCl,
0.2g of KCl, 1.44g of Na2HPO4 and 0.24g of KH2PO4 were dissolved in 80 mL H2), the
pH adjusted to 7.4, topped to 100 mL and autoclaved.
Appendix 4: Protein Purification Buffers
a) Native Purification Buffer: To prepare 100 mL 5X Native Purification Buffer
(250 mM NaH2PO4 and 2.5M NaCl), 3.5g sodium phosphate, monobasic and 14.6g
sodium chloride were dissolved in 90 mL H2O. The pH was adjusted to 8.0 and the
volume topped to 100 mL.
b) Imidazole: To prepare 100 mL 3M Imidazole, 20.6 g Imidazole, 8.77 mL
Stock Solution A (200 mM sodium phosphate, monobasic (NaH2PO4), 5 M NaCl and 1.23
mL of Stock Solution B (200 mM sodium phosphate, dibasic (Na2HPO4) and 5 M NaCl
were added to 80 mL H2O. The ph was adjusted to 6.0 and the final volume brought to
100 mL.
c) Native Binding Buffer: To prepare 30 mL native binding buffer, 30 mL 1×
Native Purification Buffer and 100 µL of 3 M Imidazole, pH 6.0 were mixed well and the
pH adjusted to 8.0.
d) Native Wash Buffer: To prepare 50 mL Native wash buffer, 50 mL of 1×
Native Purification Buffer and 335 µL mM of 3M Imidazole, pH 6.0 were mixed and the
pH adjusted to 8.0.
e) Native Elution Buffer: To prepare 15 mL, 13.75 mL of 1× native purification
buffer was mixed with 1.25 mL of 3 M Imidazole, pH 6.0. The pH was adjusted to 8.0.
100
Appendix 5. Insect cell culture media
a) Complete TNM-FH media: To prepare 100 mL, 90 mL of Grace insect media,
supplemented was mixed with 10 mL heat inactivated fetal bovine serum (FBS). 1 mL
pen-strep was added to give a final concentration of 100 U/mL penicillin and 100 µg/mL
streptomycin.
b) Freezing medium: To make 100 mL of freezing medium, 70 mL of complete
TNM-FH medium was mixed with 10 mL of heat inactivated FBS and 20 mL of 100%
glycerol.
Appendix 6. Quantities of the purified Recombinant VATs
Table A-2: Quantification results of Purified recombinant VATs expressed in
bacterial and insect cells
Recombinant protein OD562nm Concentration (µg/mL)
VSG 3 pBAD/His A 2.529 157.335
VSG 4 pBAD/His A 2.382 84.605
VSG 4 pRSET/His B (F0) 2.171 74.383
VSG 4 pRSET/His B (F1) 1.482 68.945
VSG 3 pIZ/V5-His 2.4215 120.3975
VSG 4 pIZ/V5-His 2.392 108.471
